development of oxidoreductase based electrochemical biosensors · patrícia raquel dos santos...

Patrícia Raquel dos Santos Rodrigues

Licenciada em Bioquímica

Development of oxidoreductase basedelectrochemical biosensors

Dissertação para obtenção do Grau de Mestre emBiotecnologia

Orientadores : Profa Doutora Maria Gabriela Machado deAlmeida, Investigadora associada, FCT-UNLProfa Doutora Sofia de Azeredo Pereira, Investi-gadora auxiliar, FCM-UNL

Júri:

Presidente: Prof. Doutor Rui Manuel Freitas Oliveira

Arguente: Profa Doutora Ana Pimenta da Gama da Silveira Viana Semedo,Professora Auxiliar convidada, Faculdade de Ciências da Univer-sidade de Lisboa

Vogal: Doutora Maria Gabriela Machado de Almeida, Investigadora Aux-iliar, FCT-UNL

Maio, 2013

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Development of oxidoreductase based electrochemical biosensors

Copyright c© Patrícia Raquel dos Santos Rodrigues, Faculdade de Ciências e Tecnologia,Universidade Nova de Lisboa

A Faculdade de Ciências e Tecnologia e a Universidade Nova de Lisboa têm o direito,perpétuo e sem limites geográficos, de arquivar e publicar esta dissertação através de ex-emplares impressos reproduzidos em papel ou de forma digital, ou por qualquer outromeio conhecido ou que venha a ser inventado, e de a divulgar através de repositórioscientíficos e de admitir a sua cópia e distribuição com objectivos educacionais ou de in-vestigação, não comerciais, desde que seja dado crédito ao autor e editor.

Acknowledgements

First and foremost, I’d like to thank my supervisor, Prof. M. Gabriela Almeida for all herhard-work and support, the work developed during this year wouldn’t have been thesame without her constant presence, commitment and all the opportunities she madepossible; I might not always show it but deep down I’m truly grateful for workingto/with her. But above all else I’m indebted to her for all the faith she has put in mesince the beginning.

I would like to acknowledge all the collaborators, which have contributed for thiswork and made it possible.

Dr. Michel Kranendonk, from Instituto de Higiene e Medicina Tropical, for providingthe CYP samples used during this work.

To Dra. Sofia Azeredo Pereira for all her help and know-how in the early stages of theproject.

A huge thank you to Dr. Mathieu Etienne who kindly received me in his lab in Nancyand always had an open door when I needed it.

To the ERASMUS program for enabling me to travel to France and perform such agreat part of the work in this thesis.

I can’t help mentioning my appreciation towards my office partners, Claúdia Nóbregaand Humberto Pedroso, for bearing with me even when I’m at my most despicable andcan’t stop ranting and venting for the life of me. If indeed someone deserves a place inheaven it’s the both of them for being so patient xD

Like Norma Jean would say “I’m selfish, impatient and a little insecure. I make mis-takes, I am out of control and at times hard to handle. But if you can’t handle me at myworst, then you sure as hell don’t deserve me at my best.” Butterfly you both *

A very special thank you to Célia Silveira, even though she didn’t have any obligationto listen to me, always did so with a lot of patience and helped me in any way she could.I can’t seem to find the right words to thank you enough. I know you like to think ofyourself like a cold person but I know the truth, deep down you’re one tiny softy =P

I have thank César, if it wasn’t for him I’d have been left stranded on the wrong sideof the river for a good portion of the year. For that and everything else, thank you.

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Last but not the least I’d like to thank everybody who has at some point wished mewell (if you don’t anymore I might have done something to deserve it), you probablyknow who you are better than I do ;)

Abstract

This thesis is divided in 2 sections, each describing the development of an oxidore-ductase based biosensor. In the first part human Cytochrome P450 1A2 (CYP1A2) electro-chemistry was studied, while the second is focused on the optimization of immobiliza-tion platforms and operation methods for amperometric biosensors, using cytochrome cnitrite reductase (ccNiR), (Desulfovibrio desulfuricans ATCC 27774) as a model enzyme.

The direct electrochemistry of P450s immobilized in water-based sol-gel thin filmswas described for the first time. The optimization of the film showed that only the com-bination of the inorganic matrix and the PEG400 enabled the direct electron transfer re-action and electrocatalytic activity towards oxygen. The amount of dissolved oxygenin solution revealed itself a significant feature in CYP’s electrochemistry – in anaerobicconditions, when small amounts of oxygen are added the PFeIII/II signal’s intensity in-creased, while in aerobic conditions it disappeared; probably PFeIII is not being regen-erated. However, this was not observed with the CYPOR complex, indicating that thereductase has an essential role in the CYP’s catalytic cycle completion; this was also sus-tained by the fact that only in its presence organic substrates catalysis (caffeine) occurs.

The hybrid sol-gel developed for CYP, was optimized for a nitrite biosensor. ccNiRwas successfully incorporated while promptly displaying catalytic currents. Althoughthe bioelectrode’s response decreases after day one, it was able to maintain a reasonablecatalytic activity over a time span of 2 weeks. Another electrode modification strategy,studied with ccNiR, was based on the electrophoretic deposition of macroporous assem-blies of single-walled carbon nanotubes. The macroporous structure was created as aresult of the presence of polystyrene beads co-deposited with the carbon nanotubes. Anincrease in the amount of material was correlated with a higher enzyme activity.

Finally, an oxygen scavenger system consisting of glucose oxidase, glucose, and cata-lase was employed for oxygen removal in an open electrochemical cell. The system com-pletely removed oxygen for over 1 h and was successfully applied to a ccNiR based nitritesensor.

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Keywords: biosensors, cytochrome c nitrite reductase, cytochrome P4501A2, sol-gel,carbon nanotubes, electrophoretic deposition.

Resumo

O presente trabalho encontra-se dividido em duas secções, cada um descrevendo odesenvolvimento de um biossensor baseado em oxidorredutases. Na primeira secção,foi estudada a eletroquímica do citocromo P450 1A2 (CYP1A2), enquanto que a segundase foca na otimização de plataformas de imobilização e métodos de operação para bios-sensores amperométricos, usando a redutase do nitrito multihémica (ccNiR) extraída deDesulfovibrio desulfuricans ATCC27774 como um enzima modelo.

A electroquímica direta de P450 imobilizado em filmes à base de filmes de sol-gelaquoso foi descrita pela primeira vez. A otimização da composição do material mostrouque a combinação da matriz inorgânica e o PEG400 facilitava grandemente a transferên-cia de electrões directa. A quantidade de oxigénio dissolvido na solução revelou-se umacaracterística importante em electroquímica do CYP - em condições anaeróbicas, em queas pequenas quantidades de oxigénio foram adicionados, a intensidade do sinal PFeIII/II

aumentou, enquanto que em condições aeróbicas desapareceu; provavelmente o PFeIII

não estava a ser regenerado. No entanto, o mesmo não foi observado com o complexoCYPOR, indicando que a redutase tem um papel essencial na conclusão do ciclo catalí-tico do CYP, o que também foi sustentado pelo facto de apenas na sua presença ocorrer acatálise de substratos orgânicos (cafeína).

O sol-gel híbrido desenvolvido para o CYP, foi adaptado para um biossensor de ni-trito no qual a ccNiR foi incorporada com sucesso. Embora a resposta do bioeléctrodotenha diminuido após o primeiro dia, a atividade catalítica foi mantida a um nivel razoá-vel ao longo de um intervalo de tempo de 2 semanas. Outra estratégia de modificação doeletrodo estudada com ccNiR, foi baseada na deposição eletroforética de nanotubos decarbono macroporosos. A estrutura macroporosa foi criado como resultado da presençade esferas de poliestireno co-depositadas com os nanotubos de carbono. Um aumento naquantidade de material foi correlacionado com o aumento da actividade enzimática.

Finalmente, um sistema eliminador de oxigénio consistindo em glucose oxidase, glu-cose e catalase, foi empregue para a remoção de oxigénio de uma célula electroquímica

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aberta. O sistema removeu completamente o oxigénio durante mais 1 h e foi aplicadocom êxito a um sensor de nitrito baseado na ccNiR.

Palavras-chave: biosensors, reductase do nitrito multihémica, citocromo P4501A2, trans-ferência electrónica, sol-gel, deposição electroforética.

Contents

1 Introduction 1

1.1 Electrochemical Biosensors . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

1.1.1 Importance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

1.1.2 History . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

1.1.3 Biosensor components . . . . . . . . . . . . . . . . . . . . . . . . . . 4

1.1.4 Immobilization techniques . . . . . . . . . . . . . . . . . . . . . . . 8

1.1.5 Amperometric biosensors . . . . . . . . . . . . . . . . . . . . . . . . 11

1.2 Nitrite Reductase based electrochemical biosensor . . . . . . . . . . . . . . 15

1.2.1 Nitrite assessment . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15

1.2.2 Nitrite Biosensors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

1.2.3 Nitrite reductases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18

1.3 Cytochrome P450 based biosensors . . . . . . . . . . . . . . . . . . . . . . . 24

1.3.1 Cytochromes P450 . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24

1.3.2 NADPH-cytochrome P450 reductase . . . . . . . . . . . . . . . . . . 26

1.3.3 Cytochrome P450 1A2 . . . . . . . . . . . . . . . . . . . . . . . . . . 27

1.3.4 P450 based electrochemical Biosensors . . . . . . . . . . . . . . . . . 27

1.4 Heme proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36

2 Experimental 37

2.1 Part A- Cytochrome P450 1A2 . . . . . . . . . . . . . . . . . . . . . . . . . . 37

2.1.1 Electrochemical measurements . . . . . . . . . . . . . . . . . . . . . 37

2.1.2 Mediated electrochemistry of cytochrome P450 . . . . . . . . . . . . 38

2.1.3 Direct electrochemistry . . . . . . . . . . . . . . . . . . . . . . . . . 38

2.2 Part B- Cytochrome c Nitrite Reductase . . . . . . . . . . . . . . . . . . . . 40

2.2.1 Electrochemical measurements . . . . . . . . . . . . . . . . . . . . . 40

2.2.2 Response to nitrite . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40

2.2.3 Hybrid sol-gel matrix . . . . . . . . . . . . . . . . . . . . . . . . . . 41

2.2.4 Macroporous Carbon Nanotubes . . . . . . . . . . . . . . . . . . . . 41

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2.2.5 Oxygen scavenger system . . . . . . . . . . . . . . . . . . . . . . . . 42

3 Results and discussion 453.1 Cytochrome P450 1A2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45

3.1.1 Cytochrome P4501A2 in the absence of CPR . . . . . . . . . . . . . 453.1.2 CYPOR complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65

3.2 Cytochrome c nitrite reductase . . . . . . . . . . . . . . . . . . . . . . . . . 693.2.1 Hybrid sol-gel matrix . . . . . . . . . . . . . . . . . . . . . . . . . . 703.2.2 Macroporous carbon nanotubes . . . . . . . . . . . . . . . . . . . . 743.2.3 Oxygen scavenger system . . . . . . . . . . . . . . . . . . . . . . . . 78

4 Conclusion 854.1 Cytochrome P450 electrochemistry . . . . . . . . . . . . . . . . . . . . . . . 854.2 Cytochrome c nitrite reductase electrochemistry . . . . . . . . . . . . . . . 86

5 Future work 895.1 Cytochrome P450 electrochemistry . . . . . . . . . . . . . . . . . . . . . . . 895.2 Cytochrome c nitrite reductase electrochemistry . . . . . . . . . . . . . . . 89

6 References 91

List of Figures

1.1 Clark electrode functioning. . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

1.2 Examples for biosensor components and measured analytes1. . . . . . . . 4

1.3 Key features of amperometric biosensors1. . . . . . . . . . . . . . . . . . . . 7

1.4 Sol-gel formation reactions from silica based alkoxide precursors. . . . . . 10

1.5 Schematic representations of surfactants in various forms, the headgroupsare represented by the red circles and the hydrophobic tails are in blue.A) Spherical micelles, B) hemimicelles, C) bilayers and D) multilayers onelectrode surfaces39. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

1.6 Direct ET (tunneling mechanism) from the active site of an enzyme to theelectrode surface77. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13

1.7 Schematic representations of the working principles of enzymatic biosen-sors with a reductase as biologic component: A)Mediated transduction,B) Direct transduction. (medox – mediator in the oxidized form; medred–mediator in the reduced form; enzymeox – reductase oxidized state; en-zymered – Reductase reduced state). . . . . . . . . . . . . . . . . . . . . . . 14

1.8 Three-dimensional structures of nitrite reductases. (a) Desulfovibrio vul-garis Hildenborough multiheme c nitrite reductase (NrfA4NrfH2 complex);the catalytic subunit (NrfA) is depicted in blue and the electron donor sub-unit (NrfH) in gray; heme groups are shown in dark red. (b) Spinach nitritereductase; siroheme is shown in dark red and iron-sulfur cluster in yellow.(c) Achromobacter cycloclastes copper nitrite reductase (trimer); the cop-per centres are shown in blue. (d) Pseudomonas aeruginosa cytochromecd1 nitrite reductase (dimer); heme c is depicted in dark red and heme din blue125. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18

1.9 Secondary structure of NrfHA viewed parallel to the membrane (grey rect-angle) with haems drawn as red sticks148. . . . . . . . . . . . . . . . . . . . 24

1.10 Proposed Cyt P450 Catalytic Cycle. RH: lipophilic compound in which anoxygen atom derived from O2 is introduced. . . . . . . . . . . . . . . . . . 25

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xvi LIST OF FIGURES

1.11 Model of the conformational equilibrium in CPR177. . . . . . . . . . . . . . 26

1.12 The secondary and tertiary structure of human P450 1A2 is shown in twoviews. The α-helices are colored blue, and the β-strands are colored brown.These secondary structure elements are designated A-L and 1-4, respec-tively, and are sequentially identified from the N terminus. The hemeprosthetic group is represented in sticks and is colored red. The substratebinding cavity is illustrated as a red mesh surface. . . . . . . . . . . . . . . 27

1.13 schematic representation of electron transfer in microsomal membrane.Electron transfer from P450 reductase to P450 (adapted from Hara (2000).161). 29

2.1 Surfactant structures. Cationic head group: A) CTAB, B) DDAB; C) DDM(n-dodecyl-b-D-maltoside). . . . . . . . . . . . . . . . . . . . . . . . . . . . 39

2.2 A) structure of sodium silicate; B) structure of poly(ethylene glycol). . . . 39

2.3 Scheme of the experimental device used for the electrophoretic depositionof carbon nanotubes201. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42

2.4 Electrochemical cell. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42

3.1 Schematic representations of the working principles of a CYP Bioelectrodewith mediated transduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . 46

3.2 A) Cyclic voltammograms of methyl viologen in the presence of CYP mem-brane entraped on a PG electrode, buffer solution 0.1 M MV, 0.1 M KCland tris-HCl buffer 50 mM pH 7.6. in the presence of varying caffeineconcentrations (0-42mM).Scan rate: 50mV s−1.B Icat variation with nitriteconcentration. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47

3.3 3 A) Cyclic voltammograms of methylene blue in the presence of CYP ona PG electrode, buffer solution 0.1 M MV, 0.1 M KCl and tris-HCl buffer 50mM pH 7.6. in the presence of varying caffeine concentrations (0–42mM).Scanrate: 50mV s−1.B . B) Icat variation with nitrite concentration. . . . . . . . 48

3.4 Consecutive cyclic voltammograms of CYP1A2–surfactant films casted onPG electrodes in 0.1 M KCl and tris-HCl 50 mM pH 7.6; scan rate 50 mV/s.A) Control electrode with CYP1A2 only, B) DDM C) DDAB, D) CTAB. . . 49

3.5 Variation of the cathodic peak currents on the potential with the scan rate(5-500 mV/s) (y=-2.48*10-8-4.48*10−10x; r2=0.992) . . . . . . . . . . . . . . 51

3.6 Cyclic voltammograms of CYP1A2 casted on PG electrodes, in 0.1 M KCland tris-HCl 50 mM pH 7.6 purged electrolyte; scan rate 50 mV/s, inthe presence of A only CYP; B CYP/Sodium Silicate; C CYP/Peg400; DCYP/Sol-gel(sodium silicate and PEG400). . . . . . . . . . . . . . . . . . . 53

3.7 Consecutive cyclic voltammograms of PG/CYP1A2–sol-gel film electrodein 0.1 M KCl and 50 mM tris-HCl buffer pH 7.6 purged with argon. scanrate 50 mV/s. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55

LIST OF FIGURES xvii

3.8 A) Cyclic voltammograms of PG/CYP1A2–sol-gel film electrode in 0.1 MKCl and 50 mM tris-HCl buffer pH 7.6 purged electrolyte at different scanrates; from inside to outside: 0.005, 0.01, 0.02, 0.05, 0.1, 0.2, 0.25 V.s−1,respectively. B) Variation of the cathodic peak currents on the potentialscan rate (y=-0.03-0.017x; r2=0.98). . . . . . . . . . . . . . . . . . . . . . . . 56

3.9 Consecutive cyclic voltammograms of PG/CYP1A2–sol-gel film electrodein 0.1 M KCl and 50 mM tris-HCl buffer pH 7.6 aerobic conditions. scanrate 50 mV/s. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58

3.10 Pathways for Biocatalytic Activation of Cyt P450s by Peroxides, OxygenReduction Formed Peroxide.by P450s on Electrodes. . . . . . . . . . . . . . 59

3.11 Cyclic voltammograms of CYP1A2–sol-gel film casted on PG electrode,buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6 purged with argon.In the presence of varying caffeine concentrations in the absence of oxygen.Scan rate: 50mV s−1. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60

3.12 Cyclic voltammograms of CYP1A2–sol-gel film casted on PG electrode,buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6 purged with argon.In the presence of varying non-purged caffeine volumes (stock solution 60mM). Scan rate: 50mV/s. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61

3.13 Cyclic voltammograms of CYP1A2–sol-gel film casted on PG electrode,buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6 purged with argon.Consecutive additions of non-purged water. Scan rate: 20mV s−1. . . . . . 62

3.14 A) Cyclic voltammograms of PG/CYPOR–sol-gel film electrode in 0.1 MKCl and 50 mM tris-HCl buffer pH 7.6 in a purged electrolyte at differentscan rates; from inside to outside: 0.005, 0.01, 0.02, 0.05, 0.1, 0.2, 0.25 V.s-1, respectively. B) Variation of the cathodic peak currents on the potentialscan rate (y=1.85x10-10−8.39x10−10x; r2=0.997). . . . . . . . . . . . . . . . . 66

3.15 Cyclic voltammograms of films casted on PG electrode in aerobic condi-tions, buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6. Scan rate:50mV s−1.of A) CYPOR–sol-gel B) CYP1A2–sol-gel film. a) 1st scan b) 20th

scan. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67

3.16 The effect of O2 binding on the electrochemistry of the CYPOR/sol-gelcasted on a PG electrode, purged buffer solution, 0.1 M KCl and tris-HCl50 mM pH 7.6. Consecutive additions of non-purged water. Scan rate:50mV s−1. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68

3.17 Cyclic voltammograms of CYPOR/sol-gel films casted on PG electrodein anaerobic conditions, buffer solution, 0.1 M KCl and tris-HCl 50 mMpH 7.6 purged with argon.scan rate, 50 mV/s.(black-line) without sub-strates (red-line) oxygen addition (blue and green lines) caffeine additionsincreased concentration. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69

xviii LIST OF FIGURES

3.18 Cyclic voltammograms of ccNiR immobilized on sol -gel films, buffer so-lution, 0.1 M KCl and tris-HCl 50 mM pH 7.6 purged with argon.scan rate,20 mV/s. A) ccNiR entrapped in a sodium silicate and PEG400 film B)ccNiR entrapped in a sodium silicate and PEG6000 film. . . . . . . . . . . 71

3.19 Electrochemical response of PG/ccNiR–sol-gel film(PEG6000) electrode tovarying nitrite concentrations (0-500µM) in 0.1 M KCl and 50 mM tris-HClbuffer pH 7.6 purged with argon. Scan rate, 20 mV/s. . . . . . . . . . . . . 72

3.20 Variation of Icat with nitrite concentration of ccNiR immobilized on sol-gelfilms A) ccNiR entrapped in a sodium silicate and PEG6000 film B) ccNiRentrapped in a sodium silicate and PEG400 film. . . . . . . . . . . . . . . . 73

3.21 Variation of Icat with nitrite concentration of ccNiR immobilized on sol-gelfilms A) ccNiR entrapped in a sodium silicate and PEG6000 film B) ccNiRentrapped in a sodium silicate and PEG400 film.A) Variation of Icat withnitrite concentration of ccNiR immobilized on PEG6000 sol-gel films overtime. B) Time effects on the biosensor sensitivity for nitrite determination.Sensitivity values were given by the slope of calibration curves performedperiodically throughout 80 days. Catalytic currents wer e measured at -0.8 V vs Ag/AgCl. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75

3.22 Electrochemical response of PG/CYP1A2–sol-gel film electrode to varyingnitrite concentrations (0-300mM) in 0.1 M KCl and 50 mM tris-HCl bufferpH 7.6 purged with argon. Scan rate, 20 mV/s. . . . . . . . . . . . . . . . 76

3.23 Catalytic current variation of the ccNir/CNT-PS layer as a function of ni-trite concentration. Macroporous SWCNTs deposition time. (•) 30s; (•)135s; (•) 180s; (•) 240s. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77

3.24 Scheme of the GOx-CAT scavenging system. . . . . . . . . . . . . . . . . . 783.25 Cyclic voltammograms of PGE/ccNiR/PEG at 20 mV s -1 in 10 mL of 0.1

M KCl and tris-HCl 50 mM pH 7.6; (a) GOx (12.5 µM, 15 UmL -1 ) and CAT(16.6 µM, 2 kU mL -1 ) in solution (b) GOx (12.5 µM, 15 UmL-1), CAT (16.6µM, 2 kU mL-1) and glucose (50 mM) in solution. . . . . . . . . . . . . . . 80

3.26 Cyclic voltammogram with PGE at 20 mV s -1 in 10 mL of 0.1 M KCl andtris-HCl 50 mM pH 7.6; (a) upon addition of GOx (12.5 µM, 15 UmL -1) and CAT (16.6 µM, 2 kU mL -1 ) and with the addition of glucose (50mM).(b) addition of 100 mM Nitrite. . . . . . . . . . . . . . . . . . . . . . . 81

3.27 Cyclic voltammogram with ccNiR/CAT immobilized in PGE at 20 mV s-1 in 10 mL of 0.1 M KCl and tris-HCl 50 mM pH 7.6; A) ( - ) addition ofGOx (12.5 µM, 15 UmL -1 ) and of glucose (50 mM).(-) after purging theelectrolyte solution with argon for 10 min (-) CAT (16.6 µM, 2 kU mL -1) (-) 100 mM Nitrite.B) control experiments performed in the absence ofccNiR. (Red line) increasing hydrogen peroxide concentrations (0-3mM);(black line) non purged electrolyte. . . . . . . . . . . . . . . . . . . . . . . 82

List of Tables

1.1 Historical landmarks in the development of enzyme based electrochemicalbiosensors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3

1.2 Types of receptors used in biosensorsa.35 . . . . . . . . . . . . . . . . . . . . 51.3 immobilization procedures for enzymes50. . . . . . . . . . . . . . . . . . . . 91.4 Generations of enzyme based amperometric and voltammetric biosensors

MET, mediated electron transfer; DET, direct electron transfer96. . . . . . . 151.5 Description and analytical parameters of nitrite reductase based biosen-

sors (N.A.—not applicable; N.D.—not determined; . . . . . . . . . . . . . 201.6 Summary of different electrode types and electrode modifications used to

construct CYP electrodes and biosensors165. . . . . . . . . . . . . . . . . . . 31

2.1 Mediators structure and formal reduction potential. . . . . . . . . . . . . . 382.2 Surfactant critical micelar concentrations. . . . . . . . . . . . . . . . . . . . 392.3 Compositions of sol-gel based matrices using TMOS and CTAB. . . . . . . 402.4 Compositions of sol-gel based matrices using Sodium silicate and CTAB. . 402.5 Compositions of sol-gel based matrices using Sodium silicate and PEG400. 40

3.1 Compilation of several electrochemical experiments with sol-gel/CYP1A2in anaerobic conditions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 64

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xx LIST OF TABLES

1Introduction

1.1 Electrochemical Biosensors

1.1.1 Importance

The sensitive and selective determination of a large number of compounds is of greatrelevance, and has always been a problem of the utmost importance scientific research.In the field of health-care, it is indispensable for the diagnosis of diseases. Biotechnology,too, requires the analysis of complex media. High selectivity, even in trace analysis, hasbeen achieved by the considerable progress in analytical instrumentation, as is reflectedby modern gas chromatography, high-pressure liquid chromatography, mass spectrome-try and atomic absorption spectroscopy. However, due to the high costs associated withthese powerful instrumental techniques they are only used in specialized laboratories.The development of methods highly selective and easy to handle is thus a key issue inanalysis. Whereas reliable sensors are available for the determination of physical pa-rameters, e.g. temperature, pressure, or sound energy, the qualitative and quantitativeanalysis of chemical composition remains difficult. Electrochemical sensors, such as pHelectrodes and Clark-type electrodes for oxygen measurement are widely used for thispurpose1-3. Biosensors may be the answer to these problems.

1.1.2 History

The term biosensor was devised by Karl Camman in 1977. He defined it as a chemicalsensor where the recognition system uses a biochemical mechanism. However, the IU-PAC did not agree on the implementation of the term until 1997. Whereupon, the IUPAC

1

1. INTRODUCTION 1.1. Electrochemical Biosensors

committee came up with their own definition, comprising Camman’s definition plus sev-eral parameters that must be fulfilled (see below)2.

Even though Karl Camman was the one who laid claim to the term, Leland C. Clark isconsidered by many the father of biosensors. He had developed the first bubble oxygena-tor for use in cardiac surgery. Conversely, when he came to publish his results, his articlewas refused by the editor since the oxygen tension in the blood coming out from the de-vice could not be measured. This instigated Clark to develop the oxygen electrode4, withwhich he meant to measure the reduction of oxygen with a platinum electrode in orderto determine the oxygenation of blood5. His first sensor failed because the blood compo-nents would adsorb on the electrode’s surface, which in turn distorted the signal. Lateron, Clark had the idea, which would change the history of biosensors, of using the cello-phane wrapper of a cigarette packet on his sensor, making it so that only low molecularweight substances, mainly oxygen, could reach the electrode surface and be measured.The reduction current indicated the oxygen concentration, and so the Clark electrodewas created2. Nowadays, Teflon is used as the membrane and this sensor remains a keytool in medicine and environmental monitoring. Afterwards, Clark developed the sensorfurther by entrapping concentrated glucose oxidase (GOx) with another semi-permeablemembrane in front of the electrode, which could then be used for multiple glucose mea-surements based on the monitoring of the reaction described on equation 1.16.

glucose+O2glucose oxidase−−−−−−−−→ gluconic acid+H2O2 (1.1)

The enzyme layer became an integrated part of the sensor. Lee Clark went on to coin theterm “enzyme electrode” at a meeting of the New York Academy of Sciences in 19622. Itwas this electrode arrangement that introduced the new sensor concept - the biosensor.A few years later, hydrogen peroxide reduction was detected on the electrode insteadof oxygen, due to limitations concerning this particular system, namely, oxygen fluctua-tions, substrate limitations/sensitivity, electrochemical interferences, among others7.

Clark’s system has been thoroughly studied and modified throughout the years, forseveral different purposes, one of which – an oxygen scavenging system – will be ex-plored in this thesis.

Figure 1.1: Clark electrode functioning.

2


Because of the volume of the literature regarding amperometric biosensors as well asspace limitations it is not possible to cite all contributions to the field. It was selectedrepresentative work in the field of electrochemical biosensing. General milestones andachievements relevant to biosensor research and development are listed in Table 1.1.

3


Table 1.1: Historical landmarks in the development of enzyme based electrochemicalbiosensors.

Date Event References

1962 First glucose enzyme electrode Clark, L. C. & Lyons, C.5

1973 Glucose enzyme electrode based on peroxide detection Guilbault, G. G. & Lubrano, G. J.8

1975 Launch of the first commercial glucose sensor system YSI Inc.

1977 Karl Cammann introduced the term “ biosensor ” Cammann, K.9

1977 Reversible ET of cytochrome c was obtained employing tin-doped indium

oxide electrodes and 4,4′ - bipyridiyl as a promoting monolayer on gold

electrodes.

Eddowes, M. J. & Hill, H. A. O.10

Eddowes, M. J., Hill, H. A. O. & Uosaki, K.11

1981 Oxidation of NADH at graphite electrodes is described for the first time. Huck, H. & Schmidt, H.-L12

Jaegfeldt, H. et al13

1982 Demonstration of in vivo glucose monitoring Shichiri, M. et al.14

1984 First ferrocene - mediated amperometric glucose biosensor by Cass et al.

The work led to the development of the first electronic blood glucose

measuring system which was commercialized by MediSense Inc. (later

bought by Abbott Diagnostics) in 1987.

Cass, A. E. G. et al.15

1987 Launch of the first glucose meter Medisense Inc.

1987 Electrical wiring of enzymes Degani, Y. & Heller, A.16

1988 Adam Heller and Yinon Degani introduced the electrical connection

(“wiring”) of redox centers of enzymes to electrodes through electron -

conducting redox hydrogels. This work was the basis for continuous

glucose monitoring employing subcutaneously implanted miniaturized

glucose biosensors.

Degani, Y. & Heller, A.17

Forster, R. J. & Vos, J. G.18; Csöregi, E.,

Schmidtke, D. W. & Heller19

Schmidtke, D. W.20

Wagner, J. G. et al.21

1988 Direct ET by means of immobilized enzymes was introduced Frew, J. E. et al.22–28

1997 IUPAC introduced for the first time a definition for biosensors in analogy

to the definition of chemosensors

Zayats, M. et al.29

1999 Launch of a commercial in vivo glucose sensor Minimed Inc.

2000 Introduction of a wearable noninvasive glucose monitor Cygnus Inc.

2002 Schuhmann et al. introduced the use of electrodeposition paints (EDPs) as

immobilization matrices for biosensors. Following work enabled the

incorporation of redox mediators into the polymer structure of EDPs

Kurzawa, C. et al.30–32

2007 An implanted glucose biosensor (Freestyle Navigator System) operated

for five days

Weinstein, R. L. et al.33

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1.1.3 Biosensor components

Nowadays, biosensors are characterized by the direct spatial combination of a matrix-bound biologically active substance - the so-called receptor - with a transducer compo-nent. For molecular recognition, biosensors may also be equipped with other biologicelements instead of enzymes. Besides nearly all kinds of electrodes, various other signaltransducers have been combined with the immobilized bio-material34. Examples of thesecomponents are given in Figure 1.2; this compilation helps one to understand which pa-rameters change during a biological recognition event in a biosensor. The choice of thetransduction process and transduction material is dependent on this knowledge as wellas the chemical approach to construct the sensing layer on the transducer surface.

Figure 1.2: Examples for biosensor components and measured analytes1.

The choice of the biological recognition element is the crucial decision when devel-oping a novel biosensor device. Most importantly, the bioreceptor needs to selectivelyreact with the analyte of interest. The bioreceptor needs to be stable under the opera-tion and should provide a reasonable long-term stability. In Table 1.2 it is possible to seedifferent types of receptors used in biosensors, that recognize specific species, and the

5


electrochemical measurement techniques linked to them29.

Table 1.2: Types of receptors used in biosensorsa.35

6


Enzymes are by far the most commonly used biological components in biosensors.Furthermore, electrochemical transduction is the most popular signaling method, withamperometry (measurement of electric current) the favored configuration. It is advan-tageous if the chemical structure of the enzyme allows the introduction of additionalfunctionalities for chemical modification with redox mediators, binding, or crosslinkingwith the immobilization matrix. In addition, the potential for tuning the properties of theredox enzyme by means of genetic or chemical techniques can be helpful for biosensoroptimization. An important factor, especially with respect to potential commercializa-tion, is that the redox enzyme is available at reasonable costs and effort. This work willfocus on membrane-bound enzymes as the bioreceptor, therefore they will be given spe-cial attention in this section.

The main advantages of employing enzymes in biosensor architectures are the fol-lowing:

i) They can exhibit a very high catalytic activity with a turnover on a per mole basiswhich makes them not only exceptional bioelectrocatalysts for effective signal am-plification in biosensors but also for biofuel cells. Good turnover frequencies kcat arein the range of up to, at least, 100 s-1.

ii) Typically, enzymes have a high selectivity for their substrates.

iii) The driving force - redox potential needed to achieve enzymatic biocatalysis - is oftenvery close to that of the enzyme’s cofactors. Therefore, biosensors can operate atmoderate potentials.

iv) In several cases, an improvement of the enzyme stability is found when enzymes areimmobilized on transducer surfaces36,37.

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Figure 1.3: Key features of amperometric biosensors1.

Knowing the most influential parameters on a specific biosensor is the basis to un-derstand and fine tune the performance of these devices in a rational manner. Figure 1.3summarizes several analytical features of typical biosensors; selectivity, sensitivity, ac-curacy, response and recovery times, as well as operating lifetime are some of the mostsignificant key factors.

An essential aspect of biosensor optimization is the elimination/reduction of the im-pact of interferences. In most samples there are components that either directly reactat the electrode surface or at the involved redox centers, or interfere with the biological

8


recognition reaction (e.g., inhibitors or other enzyme substrates). Additional problemssuch as leakage from the sensing layer, loss of enzyme activity or electrode fouling mayoccur. Therefore, changes in sensitivity and baseline drifts are bound to arise duringbiosensor operation. Suitable strategies are needed to ensure reproducible and quantita-tive results38. For applications aimed for the general public, it is imperative to character-ize and optimize the biosensor architecture under actual measuring conditions.

Besides dictating the efficiency of electrochemical transduction, the biosensor designalso plays a fundamental role in defining the response features. For example, sensitivityand response time, which strongly depends on mass transfer limitations, are particularlyinfluenced by the characteristics of the immobilization matrix39.

1.1.4 Immobilization techniques

The main limitation for successful applications comes from the difficulty to elaboratereagentless devices in which all components enabling the bioelectrochemical detection(i.e. enzyme(s), cofactor and electro-catalytic system for cofactor detection and regenera-tion) are immobilized in a durable and active form40. The immobilization of biomoleculespermits the re-use of costly biological molecule and allows a significant simplification ofthe analytical apparatus.

Controlled immobilization techniques are useful to more than allowing the re-use orcontinuous use of industrial enzymes. The immobilization and subsequent post-immobilizationtechniques can be very advantageous to greatly increase activity–stability properties ofenzymes41. For example, inactivation of multimeric enzymes may be strongly influencedby the dissociation of subunits. On the other hand conformational changes promotedby any denaturing agent (heat, pH, organic solvents) on the small fraction of dissociatedmonomers could be much more rapid and intense42,43.

The use of biological species such as proteins, peptides, nucleic acids and even wholecells in biosensors relies largely on the successful immobilization of the bioreceptor ina physiologically active form. Typical methods to immobilize bioreceptor onto inor-ganic, organic or polymeric surfaces have been based on physical adsorption44, cova-lent binding45, entrapment in semi-permeable membranes46 and microencapsulation intopolymer microspheres and hydrogels47,48. Notwithstanding problems such as leachingand desorption44, denaturation and difficulty controlling the orientation of the biomolecule49,these methods have proven to be amenable to the immobilization of a large number ofbiomolecules.

Various methods have been described for protein immobilization (Table 1.3). In gen-eral, biosensors using adsorbed enzymes or proteins are insensitive and unstable, exceptfor a few cases, this procedure alone is rarely used in biosensor construction.

The relative lack of long-term stability in biological molecules is the most seriouslimitation in commercializing biosensors. Under some circumstances, the immobilized

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Table 1.3: immobilization procedures for enzymes50.

preparation displays a decrease in stability. Chibata51 reports that, of 50 enzyme immo-bilizations studied, 60% showed an increase in stability, 24% were unaffected and 16%showed a decrease in stability.

As it is, in this thesis, special attention will be given to sol-gel immobilization andsurfactant films. The interest of sol–gel chemistry for bioencapsulation has been largelydiscussed in the literature52 for several types of biosensors, but most importantly it hasbeen successful in protein immobilization on electrode surfaces53,54. Sol-gel matrices areproduced from a starting colloidal suspension of sol-gel precursors –the sol – which un-dergoes hydrolysis and condensation reactions to form the gel (Figure 1.4).

The sol-gel entrapment can be a relatively gentle chemical procedure that is carriedout at room temperature so that many of the molecules can endure the entrapment. Thesol-gel methodology has been extensively used to immobilize soluble proteins showingthat the majority of them can be encapsulated with retention of their native structure andfunctionality and an enhanced stability52,55–58.

Bioencapsulation within these materials is usually obtained from the hydrolysis ofalkoxide precursors (usually tetraethyl orthosilicate, TEOS, or tetrametyl orthosilicate,TMOS) resulting in a colloidal sol solution. Subsequently, a buffered aqueous solutioncontaining the biomolecule of interest is added to the sol producing a polycondensationreaction that leads to the formation of a transparent highly porous gel that encloses thespecies within its pores. The popularity of sol-gel materials can be attributed to this andother significant factors, namely54:

(a) The ability to generate an almost infinite number of organic- inorganic hybrids that

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display both the mechanical stability of a rigid inorganic framework and the particu-lar reactivity (e.g., selective recognition, optical properties) of the organic component;

(b) The fact that sol-gel-derived materials can be used to encapsulate biomolecules (e.g.,enzymes, antibodies, or other proteins) in a functional state;

(c) The discovery of the supramolecular template approach, which can generate orderedmesostructures over long length scales.

Figure 1.4: Sol-gel formation reactions from silica based alkoxide precursors.

Surfactants are surface active agents composed of a charged or polar head group anda hydrophobic nonpolar tail. The tail is generally a hydrocarbon chain with 6-22 carbonatoms. The hydrophilic region of the molecule – the head group – may be positive, neg-ative, neutral or zwitterionic, thus categorizing the surfactant as cationic, anionic, non-ionic or zwitterionic59,60. Surfactants with longer hydrocarbon chains are also less proneto denature proteins than short chained compounds. Zwittergents like amphoteric sulfo-betaines are usually more inactivating than non-ionic reagents59,60. Surfactant moleculescan be adsorbed at the interface between two bulk phases such as the electrode/solu-tion interface; lowering the interfacial tension between the two and facilitate the contact.Therefore, these surface active agents are capable of modifying and controlling electrodesurface properties61. Surfactant molecules can aggregate to form supra-molecular struc-tures with specific regions of hydrophilic and hydrophobic character. In aqueous solu-tion, above the critical micellar concentration (CMC), they form spherical micelles withthe hydrophilic head groups facing the solution and the hydrocarbon chains orientedtowards the interior of the structure Figure 1.5 59. The CMC is characteristic of each sur-factant and in addition to micelle formation it is also correlated with abrupt changes orsharp discontinuities in the physical properties of the surfactant, such as conductivityand surface tension62,63.

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Figure 1.5: Schematic representations of surfactants in various forms, the headgroups arerepresented by the red circles and the hydrophobic tails are in blue. A) Spherical micelles,B) hemimicelles, C) bilayers and D) multilayers on electrode surfaces39.

Electrochemistry mainly takes advantage of two properties associated with these sur-face active agents: their ability to adsorb at interfaces and to generate membrane-likestructures61. In particular the membrane bound proteins incorporated in surfactant filmsare expected to experience a more natural environment, closer to the physiological sur-roundings they are taken from. Accordingly, surfactant films can help to promote andenhance protein electrochemistry and catalysis. Obvious cases are membrane-boundproteins62, such as the enzyme employed in this thesis. Various soluble proteins havealso shown enhanced electron transfer in surfactant films61,64-66. Myoglobin and somecytochrome P450 isoforms, for example, have showed good reversible electrochemistryin surfactant films of DDAB (Dodecyldimethylammonium bromide) at PG electrodes, asopposed to the bare ones, where no DET was observed. According to spectroscopic stud-ies, both proteins were also able to keep their native conformations while embedded inthe films62,63,67.

The good performance of proteins in surfactant films has been associated to the strongadsorption of the surfactant at the electrode-film interface, therefore avoiding the de-naturative adsorption of proteins or of other macromolecules that could block electrontransfer63. Also, the well-defined microstructure and dynamics of the surfactant films al-lows good diffusion within the layer. These films can also help to guide the orientation ofimmobilized proteins. Which may not be related with the surfactant headgroup chargebut with the interaction between the protein and the hydrophobic bilayers68.

Rusling has pioneered the use of surfactant films on carbon surfaces for direct electro-chemistry of heme proteins69-71. The surfactant is deposited onto the electrode, resultingin the formation of bilayers and micelles into which the protein is incorporated. The endresult is a system that supports rapid and reversible ET between the electrode and theenzyme; typically, standard rate constants between 50 and 300 s-1 are observed. Protein-surfactant film voltammetry is now a routine method for studying the redox chemistry of

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Heme transfer proteins: flavocytochromes72, engineered mutants73, as well as bacterial74

and mammalian75 variants, have been investigated.

1.1.5 Amperometric biosensors

As previously mentioned, a biosensor is generally defined as a sensing device consistingof a biological recognition element in intimate contact with a suitable transducer which isable to convert the biological recognition reaction, or eventually the biocatalytic process,into a measurable electronic signal. In the particular case of an amperometric sensor,the redox species in the sensing layer are oxidized or reduced at the transducer surfacegenerating a current through the electrode. Assuming an enzyme catalyzed oxidation ofa substrate (e.g. glucose, lactate, alcohol, etc.), either a prosthetic group integrated withinthe enzyme (e.g. FAD, PQQ, heme, transition metals) or a co-substrate (e.g. FMN, NAD+,NADP+) has to be reduced, intermediately storing the transferred redox equivalents. Ameasurable current through the electrode is related to the re-oxidation of the prostheticgroup or the co-substrate in order to regenerate the enzyme and make it available forfurther substrate recognition and conversion reactions. Hence, a signal is only obtainedif the transfer of electrons between the intermediately reduced enzyme (i.e. the prostheticgroup or the co-substrate) and the electrode is possible. From this discussion it is evidentthat the specific features of the biosensor are highly dependent on the kinetics of thiselectron transfer process. Consequently, an essential prerequisite for the developmentof amperometric biosensors with high sensitivity and fast response characteristics is toestablish a fast electron transfer (ET) from the biological component to the electrode.

1.1.5.1 Electron transfer

At a first glance, the easiest ET mechanism in an amperometric biosensor would be thedirect electrochemical recycling of the enzyme’s prosthetic group at the electrode sur-face involving electron tunneling mechanisms (Figure 1.6). However, according to theMarcus theory76, the ET kinetics between two redox species is determined by the drivingforce (i.e. the potential difference), the reorganization energy (which qualitatively reflectsthe structural rigidity of the redox species) and the distance between the two redox cen-ters. Obviously, the ET distance between the prosthetic group and the electrode surfaceis rather long due to the shielding by the protein shell, and direct ET via a tunnelingmechanism is therefore rarely encountered.

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Figure 1.6: Direct ET (tunneling mechanism) from the active site of an enzyme to theelectrode surface77.

Thus, the main aim in the design of optimized amperometric biosensors is to en-sure fast ET processes based on electrode architectures with predefined ET pathwaysinterconnecting the redox site within the enzyme and the electrode surface. The elec-tron transfer in biology usually involves initial protein-protein complex formation basedon the complementarity of the docking sites. Efficient protein-electrode reactions ap-pear to have some similarities to the way in which proteins act with their natural redoxpartner78. Therefore, methods for chemically modifying electrode surfaces as to mimicthe biological situation were developed. The heterogeneous electron transfer betweenproteins and electrodes may be coupled with other reactions where the proteins act asvectorial mediators79,80.

The principle of direct electrochemistry of redox proteins, can be explained by com-paring this technique to the more traditional solution essays of enzyme activity. In homo-geneous enzyme kinetics, the enzyme may be mixed in a cuvette with its substrate anda redox partner (otherwise called a mediator or co-substrate), providing a source/sink ofelectrons for the redox transformation of the substrate and whose absorbance dependson its redox state. In the steady state, the rates of substrate and co-substrate transforma-tions are equal to the turnover rate of the enzyme and can therefore be determined byfollowing the absorbance change of the solution81.

In direct electron transfer (DET) the increase in catalytic currents is a result of the en-zyme’s cofactors direct regeneration (Figure 1.7A). This type of communication with theelectrode, is typically harder to achieve due to the insulating nature of protein polypep-tide chains and the location of redox centers, which are commonly deeply buried withinthe protein structure. Furthermore, the protein molecules can have an unfavorable ori-entation on the transducer surface, sometimes contributing to an increased distance be-tween the electroactive centers and the electron transfer partner (the electrode) and thushindering the electron exchange39.

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Figure 1.7: Schematic representations of the working principles of enzymatic biosensorswith a reductase as biologic component: A)Mediated transduction, B) Direct transduc-tion. (medox – mediator in the oxidized form; medred –mediator in the reduced form;enzymeox – reductase oxidized state; enzymered – Reductase reduced state).

Nowadays is still common to think that oxireductases are too large and too fragile tointeract directly with a solid electrode without being at least partly denaturated. It is usu-ally held that because the active site of these enzymes is deeply buried in the protectiveprotein matrix, direct electron exchange at an electrode can only occur under exceptionalconditions. However, more than 20 years have passed since it was shown that directelectron transfer (DET) can occur between an electrode and a large, catalytically activeenzyme,82,93 and about one hundred examples have already been reported81,94.

Unlike in DET, the use of redox mediators takes control over the protein reaction cen-ter away from the electrode95(Figure 1.7B). On the one hand, this can be detrimental to theselectivity of detection, because the mediator can react with other species present, and,for biosensors, can require a more complex manufacturing process, additional reagents,and sophisticated immobilization methods. On the other hand, redox mediation canovercome the frequently sluggish electron communication of enzyme redox centers withelectrodes, thus increasing ET rates96. Mediated transfer of redox equivalents is the work-ing principle of second-generation amperometric biosensors. Mediators are character-ized by having a high heterogeneous ET rate that does not compromise electrochemicalreversibility and, at the same time, homogeneous, rapid electron exchange with the en-zyme. Both oxidized and reduced forms of the mediator should be stable and unreactivewith oxygen; also, the reaction should not depend on pH. Furthermore, when select-ing a redox mediator for biosensor applications it is important to consider toxicity, bio-compatibility, ease of immobilization, and, very importantly, the redox potentials. Lowoperating potentials are preferred, enabling appropriate enzyme reaction transductionwhile avoiding side reactions. In other words, a mediator should shift redox potentialsfrom the extreme values necessary to detect target analytes, to values near zero, at which

15

1. INTRODUCTION 1.2. Nitrite Reductase based electrochemical biosensor

fewer interfering species are reduced or oxidized95,97,98. In mediated electrochemistry,the consumption of the redox partner is detected as a current wave resulting from itselectrochemical recycling on the electrode; only the mediator interacts with the electrode,and the homogeneous catalytic process which occurs in the bulk of the electrochemicalcell is fundamentally the same as that in solution assays81.

Biosensors can be grouped into generations according to the modes of signal trans-fer between a redox enzyme and an electrode, i.e., via the natural secondary substratesor products of the enzyme reaction (first generation), via artificial (either synthetic orbiological) electron mediators (second generation) or via direct electronic contact (thirdgeneration)34,99,100. A typical first generation biosensor is a glucose sensor with gel en-trapped glucose oxidase on a Clark-type electrode101,102 and also many variants of cou-pled enzymes on oxygen-sensitive electrodes103.

Table 1.4: Generations of enzyme based amperometric and voltammetric biosensors MET,mediated electron transfer; DET, direct electron transfer96.

1.2 Nitrite Reductase based electrochemical biosensor

1.2.1 Nitrite assessment

Nitrate (NO−3 ) and nitrite (NO−

2 ) are rarely found without each other, because their chemistriesare practically indissociable.

Nitrate occurs naturally in soils containing nitrogen-fixing bacteria, decaying plants,septic system effluents, and animal manure. Among the artificial sources of nitrate thereare nitrogenous fertilizers and airborne nitrogen compounds emitted by industry andmotor vehicles. Nitrate penetrates through the soil and remains in groundwater fordecades104; groundwater is the source for >50% of drinking water supplies, 96% of pri-vate water supplies, and an estimated 39% of public water supplies105.

Water containing a high concentration of nitrite can create serious problems, for ex-ample, eutrophication of aquatic systems and potential hazards to human health106. Theeutrophication in rivers, lakes, and coastal waters has become one of the most prevalent

16


environmental problems in recent years106,107. The health implications of exposure to ni-trate in drinking water were first reported in 1945 by Comly after observing cyanosis ininfants in Iowa, where well water was used in formula preparation108. These implica-tions made it so that nitrite has been considered an important toxic agent109. However,as of late, there have been many new findings that imply the nitrite’s role in the humanorganism may not be so straightforward, the main controversy is that nitrite’s ingestionmight be beneficial for human health despite it’s being considered solely hazardous sincethe 1940’s110-122.

In order to manage environmental and health risks, deriving from exposure to theseions, governmental agencies have implemented rules and directives to restrict the levelsof NO−

3 and NO−2 in drinking waters and food products. European directive 98/83/EC

has established the maximum admissible levels of nitrate and nitrite in drinking waterat 50 and 0.1 ppm, respectively. Likewise, the World Health Organization has set theselimits at 50 ppm (NO−

3 ) and 3 ppm (NO−2 ) (WHO/SDE/WSH/07.01/16). More recently,

following the European Food Safety Authority recommendations, 2006/52/EC directivehas reduced the authorized levels for these ions in meat and other food products, whichshould be controlled on the basis of added rather than residual amounts (e.g. 150 mg/kgof nitrites in meat products). Furthermore, the determination of nitrite in human physio-logical fluids is also commonly used for clinical diagnosis. As a result, there is a growingdemand to detect nitrite in food, drinking water and environmental samples123.

1.2.2 Nitrite Biosensors

In the last decades, biosensor technology has been exploited as a route to provide reliablenitrite quantification in complex samples. Hence, several protein electrodes and opticaldevices are described in the literature2,34,39,124-130.

When it comes to nitrite biosensors, the bulk of the approaches make use of mediatedelectron transfer by employing redox mediators (e.g. viologen derivates) that display afast and reversible electrochemical response and are able to shuttle electrons rapidly tothe redox centers of the proteins39.

As transducing modes one could find a vast predominance of electrochemical ap-proaches, the largest group being the voltammetric and/or amperometric ones and asmall number being based on potentiometric or conductimetric platforms. Alternatively,the spectroscopic changes that take place during the catalytic cycle were also employedin the construction of optical biosensors. The strategies proposed for protein immobiliza-tion have relied on a variety of materials, ranging from non-conducting polymers, elec-tropolymerized films, redox active clays, sol-gel silica glasses, carbon nanotubes, metalnanoparticles and DNA tethers, either alone or in composite formulations. Although theconstruction of these sensing devices is far from trivial, major progresses have been madeover the last decade. After the preliminary studies carried out using non-immobilizedelectron carrier species, fully integrated biosensors based on mediated electrochemistry

17


have become a common configuration. More advanced strategies operating in the un-mediated mode via DET131 and exploiting nanostructured materials as electrodes inter-faces were recently proposed35. In parallel, stability has been substantially improvedthrough the construction of leak free devices and the use of protecting coats. Very re-cently, the screen printing technology was successfully employed, opening up the routefor miniaturization132.

The less used, mediatorless approaches - the ones on which this thesis will focus -are based on the direct electron transfer between the nitrite reductases and the electrodematerial.

Table 1.5 summarizes some of the most recent and representative works done in thefield. A detailed analysis of this table clearly demonstrates that although electrode mod-ifications are made with all sorts of materials, combined with polymers, silicates, surfac-tants or ionic liquids, the systems share many common features. Direct electron transferbetween proteins and electrodes is frequently promoted by the modifying matrix and theintensity of the catalytic currents is correlated with nitrite concentration; amperometrictransduction is a frequent option; and the working electrodes are always made of carbonmaterials (usually, glassy carbon (GC)).

Overall, the tabulated analytical parameters were highly variable. For instance, thedetection limit can be as low as 0.06 µM and go up to 700 µM. Though, the substratepromiscuity of all these heme containing proteins puts in risk the selectivity of determi-nation, so they are not recommended for the construction of a selective nitrite biosensor.Instead, much more selective biocatalysts should be employed. The next section aims todescribe all nitrite biosensing systems reported hitherto that make use of highly selectiveenzymes for nitrite reduction.

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1.2.3 Nitrite reductases

Nitrite reducing enzymes (NiRs) are the natural candidates for playing the role of biorecog-nition element in the nitrite biosensing devices. Four classes of NiRs have been recog-nized so far (Figure 1.8), all of which have already been used in biosensor applications.They can be grouped according to the type of co-factors and reaction product133,134.

Figure 1.8: Three-dimensional structures of nitrite reductases. (a) Desulfovibrio vulgarisHildenborough multiheme c nitrite reductase (NrfA4NrfH2 complex); the catalytic sub-unit (NrfA) is depicted in blue and the electron donor subunit (NrfH) in gray; hemegroups are shown in dark red. (b) Spinach nitrite reductase; siroheme is shown in darkred and iron-sulfur cluster in yellow. (c) Achromobacter cycloclastes copper nitrite re-ductase (trimer); the copper centres are shown in blue. (d) Pseudomonas aeruginosacytochrome cd1 nitrite reductase (dimer); heme c is depicted in dark red and heme d inblue125.

There are two types of ammonia forming nitrite reductases: cytochrome c nitrite re-ductases (ccNiRs), which are multi-heme enzymes isolated from sulfate or sulfur reduc-ing bacteria, and sirohemic nitrite reductases, which contain a siroheme and an iron-sulfur cluster and are commonly purified from photosynthetic organisms such as plants,algae and cyanobacteria. This group of enzymes is able to catalyze the six electron reduc-tion of nitrite to ammonia, according to the following equation:

NO−2 + 8H+ + 6e− → NH+

4 + 2H2O (1.2)

The nitric oxide forming nitrite reductases are dissimilatory NiRs involved in bacte-rial denitrification; they catalyze the one electron reduction of nitrite to nitric oxide.

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NO−2 + 2H+ + e− → NO +H2O (1.3)

There are two different types: the copper-containing nitrite reductases (CuNiRs), withtype-I and type- II copper centers, and the cytochrome cd1 nitrite reductases (cd1NiRs)that comprise a c-type and a d1-type heme134.

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21 | P a g e

Table 1-5 Description and analytical parameters of nitrite reductase based biosensors (N.A. —not applicable; N.D.—not determined;

Enzyme Source Sensor preparation Transducing mode Electron transfer Linear Range Detection

Limit Sensitivity Reference

Sirohemic

NiR Spinach leafs enzyme + BSA + glutaraldehyde Potentiometric N.A. 0.1–50 mM N.D. N.D. 135

ccNiR D. desulfuricans GC/casting of enzyme + polyacrylamide

(mediator in solution)

Voltammetric

MET

(methyl viologen) up to 200 µM N.D. N.D.

136

ccNiR D. desulfuricans GC/casting of enzyme + polyacrilamide DET up to 200 µM N.D. N.D.

ccNiR D. desulfuricans

GC/dispersion of poly(pyrrole-viologen)

+ enzyme mixture followed by

electropolymerization

Voltammetric

MET

(poly(pyrrole-

viologen))

5.4–43.4 µM 5.4 µM 1,721 mA M−1cm−2 128


GC/casting of Nafion + enzyme/

incorporation of

mediator

Voltammetric MET

(methyl viologen) 75–800 µM 60 µM 445 mA M−1cm−2 130


GC/casting of [ZnCr-AQS] LHD +

enzyme/ glutaraldehyde vapor cross-

linking

Amperometric MET

(AQS)

0.015–2.350

µM 4 nM 1,824 mA M−1cm−2 132


gold/casting of Nafion + enzyme +

mediator + glycerol + BSA/

glutaraldehyde vapor cross-linking

Conductimetric MET

(methyl viologen) 0.2–120 µM 0.05 µM 0.194 µS/µM 137

ccNiR D. desulfuricans pyrolytic graphite/casting of EETMS sol/

casting of enzyme Amperometric DET 0.25–50 µM 120 nM 430 mA M−1cm−2 124

22 | P a g e

ccNiR D. desulfuricans graphite/casting of SWCNTs

dispersion/casting of enzyme Voltammetric DET up to 150 µM N.D. 2,400 mA M−1cm−2 138

ccNiR S. deleyianum

graphite and mediator composite/casting

of enzyme + poly(carbamoyl sulfonate)

hydrogel membrane

Amperometric MET (phenosafranin) up to 250 µM 1 µM 446.5 mA M−1cm−2 139

cd1NiR P. denitrificans

graphite/enzyme entrapment through

dialysis membrane


Amperometric MET

(1-methoxy PMS)

4.35–65.2

µM* N.D. N.D. 140

cd1NiR P. denitrificans

graphite/enzyme entrapment with dialysis

membrane


Amperometric MET

(1-methoxy PMS) up to 750 µM 10 µM 33 mA M−1cm−2 139

cd1NiR P. pantotrophus enzyme incorporated in bulk sol-gel

monoliths of TEOS Optical N.A.

0.075–1.250

µM 0.075 µM N.D. 141

cd1NiR P. pantotrophus enzyme in controlled pore glass beads of

isothiocyanate Optical N.A. 0–4 mM 0.93 µM 19.5 nM−1 142

cd1NiR M. hydrocarbono-

clausticus

graphite/casting of polyvinyl alcohol +

enzyme + mediator followed by

photopolymerization

Amperometric MET

(cyt-c552) 10–200 µM 7 µM 2.49 A cm2 µM−1 129

CuNiR R. sphaeroides GC/electropolimerization of PPB/ casting

of enzyme + PBV Voltammetric MET (PPB) up to 50 µM 1 µM 789 mA M−1cm−2* 143

CuNiR R. sphaeroides

GC/casting of poly(vinyl alcohol) +

mediator + enzyme/ casting of

poly(allylamine hydrochloride)/

casting of hydrophilic polyurethane

Amperometric MET

(methyl viologen) 1.5– 1.5 µM 170 mA M−1cm−2* 144

23 | P a g e

CuNiR A. faecalis

gold/enzyme entrapped with dialysis

membrane


Amperometric MET

(1-methoxy PMS) 0–22 µM* 0.22 µM* N.D. 145

CuNiR A. faecalis

gold/dip-coating in (cysteine) thiolated

hexapeptide

(enzyme and mediator in solution) Voltammetric

MET (pseudoazurine) 200–1,500

µM N.D. N.D.

146 gold/dip-coating in (cysteine) thiolated

hexapeptide

(enzyme and mediator in solution)

MET (ruthenium

hexamine) 1–100 µM N.D. N.D.


1.2.3.1 Cytochrome c Nitrite Reductase

The cytochrome c nitrite reductase from D. desulfuricans was used as biorecognition ele-ment, in the experiments addressed in this thesis. This enzyme is involved in the path-way of dissimilatory nitrite reduction to ammonia thereby playing a crucial role in thebiogeochemichal nitrogen cycle.

As mentioned above, ccNiR catalyzes the six-electron reduction of nitrite to ammonia(NH+

4 ) using electrons from the oxidation of formate or hydrogen, mediated through amenaquinone and a quinol- oxydizing system147.

ccNiR also catalyzes the reduction of nitrite oxide and hydroxylamine to ammoniaand of sulfite to sulfide It is usually found as a membrane associated complex with atransmembrane subunit. The physiological form of the enzyme is believed to be a dou-ble trimer of 2 NrfA and 1 NrfH subunits148 (Figure 1.9) in vitro, the protein complexesassociate each other forming large aggregates (min. 890 kDa)149.

Periplasm

Cytoplasm

Figure 1.9: Secondary structure of NrfHA viewed parallel to the membrane (grey rectan-gle) with haems drawn as red sticks148.

The catalytic subunit NrfA (61 kDa), which is associated to the periplasmic mem-brane, is a pentaheme cytochrome c-type where the short distances between hemes allowa fast and efficient electron transfer150,151. It can be assessed by two channels that reachthe protein surface.

They were proposed to be the substrate and product channels because they have anoverall electrostatic charge opposed to the anion NO−

2 substrate and cation NH+4 product

that should flow through them. This is thought to contribute to the high catalytic activityof this enzyme151-153 . The product channel in D.desulfuricans ccNiR is partially blocked

24


by a polypeptide segment; accordingly, the product may have to find a different routeto exit the active site or a conformational change may allow this to occur through thedesignated product channel151. NrfH (19 kDa) is a small membrane-bound cytochromecomprising four c-type heme groups and it serves a double purpose; it anchors the cat-alytic subunits to the membrane and serves as a quinol oxidase, transferring electronsfrom the quinone pool to the catalytic subunits148,149. It is composed of a transmembranehelix and a globular hydrophilic domain that houses the four hemes. The NrfH subunitinteracts with the NrfA dimmer in an asymmetrical way, with only one of the catalyticmonomers receiving electrons directly from NrfH.

25

1. INTRODUCTION 1.3. Cytochrome P450 based biosensors

1.3 Cytochrome P450 based biosensors

1.3.1 Cytochromes P450

he cytochromes P450b (CYP) are a ubiquitous superfamily of mixed function oxidases.They play an important role in the detoxication of bioactive compounds and hydrophilicxenobiotics being formed inside cells in living organisms (cholesterol, saturated andunsaturated fatty acids, steroids, prostaglandins and others) or from external souces(medicines, drugs, food supplements, and environmental pollutants)166,167. These en-zymes represent a superfamily of b-type hemeproteins with a catalytic activity towardstwo substrates: oxygen and organic substances The general monooxigenase reaction bywhich the CYPs metabolize their substrates is the following.

RH +O2 +NA(P )H +H+ → ROH +H2O +NAD(P )H+ (1.4)

The generic reaction catalyzed by P450 (Figure 1.11) implies the reduction of molecu-lar oxygen with two electrons that are supplied by various redox partners168.

Figure 1.10: Proposed Cyt P450 Catalytic Cycle. RH: lipophilic compound in which anoxygen atom derived from O2 is introduced.

26


These enzymes are unique in being able to hydroxylate non activated carbon atoms(C-H bonds)156,166,167. The products of substrates’ metabolism usually serve as regu-lators in cells or are excreted from organisms. Cytochromes P450 are capable of metab-olizing over 1,000,000 chemicals and involve about 60 distinct classes of biotransforma-tion reactions, e.g. hydroxylation, N-, O- or S-demethylation, dealkylation, epoxydationamong others169,170. This particular feature of P450s makes them one of the most studiedproteins as they are highly promising for use in farmaceutical drug assays and in stereo-directed synthesis of steroids171. For eukaryotic microsomal P450, the membrane-boundform of NADPH cytochrome P450 reductase (CPR) is the primary source of electrons168.Electrons from NAD(P)H flavins ferric form of cytochrome P450 (Fe3+). Resting P450sare in the ferric form (Fe+3 ), one electron reduction of ferric form leads to ferrous state(Fe+2 ) of hemoprotein, which can bind oxygen172. During reduction of P450s according tothe scheme Fe+3 +e− → Fe+2 NADPH or NADH are exhausted159. The P450s share a com-mon fold that is unique to this enzyme class. P450s are mainly β-helical, with the hemecofactor sandwiched between a larger helix-rich (alpha) domain and a small β-sheet-rich(beta) domain. A core structure around the heme cofactor provides the scaffold that al-lows oxygen activation by the P450s. The heme iron is equatorially coordinated by fourpyrrole nitrogens from the heme b macrocycle, and axially coordinated by a conservedcysteine as the proximal ligand and typically by a water molecule as the distal ligand (inthe P450 resting state).

1.3.2 NADPH-cytochrome P450 reductase

As previously mentioned, for eukaryotic microsomal P450s, the membrane bound formof cytochrome P450 reductase (CPR) orchestrates the stepwise electron transfer fromNADPH to the cytochrome P450 heme center168,173,174. NADPH-cytochrome P450 oxi-doreductase transfers electrons from NADPH to cytochrome P450 and catalyzes the one-electron reduction of many drugs and foreign compounds. CPR is a 78 kDa membraneanchored multidomain enzyme composed of a FMN-containing flavodoxin like domainassociated with a FAD-containing ferrodoxin reductase like domain175 probably origi-nating from horizontal gene transfer or exon fusion. A folded connecting domain joinsthe two sections. By using its two flavin cofactors and a peculiar electron transfer cycle,CPR is able to split the dielectronic flux of NADPH by sequential electron transfer toexternal acceptors176.

Electron transfer in biological systems occurs in the presence of electron donors suchas NADPH and NADP, and is generally affected by electron transfer mediators, suchas flavin nucleotides. In the presence of substrate and dioxygen, the monooxygenationreaction of the CPR-CYP enzyme system is dependent on the binding of NADPH to CPR.Electrons, in the form of a hydride anion are transferred from NADPH to FAD. ReducedFAD then transfers single electrons to FMN, which in turn reduces the prostetic hemeiron of the CYP178,179.

27


Figure 1.11: Model of the conformational equilibrium in CPR177.

1.3.3 Cytochrome P450 1A2

Human CYP1A2 is one of the major CYPs in human liver; it metabolizes a variety of clin-ically important drugs (e.g., clozapine, tacrine, tizanidine, and theophylline), a numberof procarcinogens (e.g. benzo[a]pyrene and aflatoxin B1), and several important endoge-nous compounds (e.g. steroids and arachidonic acids). Like many other CYPs, CYP1A2is subject to induction and inhibition by a number of compounds, which may providean explanation for some drug interactions observed in clinical practice. A large inter-individual variability has been observed in the expression and activity and in the elimi-nation of drugs that are mainly metabolized by CYP1A2. This is largely caused by genetic(e.g., Single-nucleotide polymorphisms) and epigenetic (e.g., DNA methylation) and en-vironmental factors (e.g., smoking and comedication). CYP1A2 is primarily regulated bythe aromatic hydrocarbon receptor (AhR). It is induced through AhR-mediated transacti-vation following ligand binding and nuclear translocation. To date, more than 15 variantalleles and a series of subvariants of the CYP1A2 gene have been identified and some ofthem have been associated with altered drug clearance and response to drug therapy180.

1.3.4 P450 based electrochemical Biosensors

Investigation of the catalytic activity of isolated cytochromes from the P450 superfamilyrequires the obligatory presence of redox partners and electron donors (NADPH) (seesection 1.3.3). However, redox partners are not obligatory for the electrochemical reduc-tion of P450 family hemoproteins, so the catalytic system is essentially simplified. Theelectrochemical approach is especially important in the case of unknown physiologicalpartners (e.g. CYP51 MT, systematic name CYP51b1)154. Electrochemical systems exe-cute a dual function: substitute partner proteins and serve as a source of electrons forredox enzymes. Electrochemistry permits to study the whole catalytic cycle of P450s

28


Figure 1.12: The secondary and tertiary structure of human P450 1A2 is shown in twoviews. The α-helices are colored blue, and the β-strands are colored brown. These sec-ondary structure elements are designated A-L and 1-4, respectively, and are sequentiallyidentified from the N terminus. The heme prosthetic group is represented in sticks andis colored red. The substrate binding cavity is illustrated as a red mesh surface.

from different viewpoints. On the one hand, the enzyme P450-electrodes are power toolsfor investigation of the catalytic properties of P450s towards new chemicals (i.e. for thesearch of new drugs with the properties of substrates or inhibitors). On the other hand,the stoichiometry of the P450’s catalytic cycle and the thermodynamics of electrochem-ical reduction of P450s can also be assessed. Additionally, electrochemistry can allowthe investigation of the peroxide shunt pathway with the help of bifunctional electrodes,therefore permitting the direct registration of hydrogen peroxide consumption by thehemeprotein155.

As mentioned above P450 enzyme electrodes aim to eliminate the complex electrontransfer machinery of the P450-reductase required in nature. For their successful elec-trochemical response two points must be addressed. First of all, the enzyme should bein its native conformational state and retain its catalytic activity after the adsorption orimmobilization procedure, i.e. it should maintain its active P450 form and not convertto the inactive P420 one. The P420 form results from the weakening or distortion of thethiolate bond of the cysteine residue that is the fifth ligand of the heme156. This bondweakening can occur upon immobilization on electrode surfaces. Secondly, care must betaken on the coupling of the electron consumption that must be linked to the conversionof the substrate into its product(s). The fine-tuning of electron delivery and proton flow iscrucial for the efficient conversion of substrate to product by cytochrome P450 enzymes.Ideally all the electrons provided by the electrode should be used in the formation ofthe product and not wasted on the production of oxygen reactive species in uncoupledreactions. Unfortunately this phenomenon does occur and it is well-documented for the

29


human drug metabolizing P450 enzymes157: oxygen and NAD(P)H are largely used bythese enzymes to produce hydrogen peroxide and superoxide anion radical or water.

The relevance of such an approach is obvious: cytochrome P450 and P450-based en-zyme electrodes may be used as biosensors in patient-tailored (personalized) medicine,high- throughput screening and drug interference studies.The biotechnological interestrelated to these enzymes in the development of biocatalysts or biosensors158-160 is dueto their versatility in the recognition of a vast array of compounds, many of which arerelevant to the development of fine chemicals, new drugs and drug products and evenenvironmentally important compounds (detected or degraded through bioremediation).However, in spite of the huge biotechnological interest, their use in bioelectrochemicaldevices has proven to be highly challenging.

The progression of P450-electrode systems into amperometric biosensors is largelydependent on the efficiency of the electrode-driven P450 activity and therefore the im-mobilization. Which is why a large part of this work is centered on immobilizationstrategies. However, there are two technical difficulties in handling P450s based elec-trochemical biosensors. The first problem is the complexity in the measurement of theirenzymatic activity.

Most P450s need another redox protein as an electron donor for their activity be-sides the lipophilic substrate, O2 and NADPH or NADH. Pathways of electron transferto P450s are shown in Figure 1.10.

NADPH

FMNHeme

ROHRH

FADO2

P450 P450 reductase

Figure 1.13: schematic representation of electron transfer in microsomal membrane. Elec-tron transfer from P450 reductase to P450 (adapted from Hara (2000).161).

For example, P450 reductase transfers electron to microsomal type P450 as shown inthe scheme in Figure 1.10. Usually, two electrons are supplied from NADPH to P450

30


through P450 reductase containing both FAD and FMN as shown in the upper part ofthis scheme.

Kinetic parameters such as Michaelis constant (KM) of the substrate highly dependon the microenvironment of the lipid membrane on which the P450 is bound. Both iso-lated P450 and P450 reductase should be coreconstituted into the same liposomes forexpressing its full enzymatic activity.

The second problem is the stability of the enzyme. Many P450s, especially membrane-bound enzymes from microsomes or mitochondria are labile. Some can be partly stabi-lized in the presence of glycerol161.

The types of electrode modifications used in CYP biosensor studies vary, as shownin Table 1.6, ranging from thin films with opposite charge adsorbed onto graphite toconductive polymers deposited onto noble metals such as gold or platinum. Most ofthese studies rely on a combination of electrochemical and analytical techniques to showthat the CYP biosensor is electroactive and can detect different drug compounds viaelectrochemically-driven catalysis. Nevertheless, a number of recent studies character-izing the active site structure and redox potential of CYPs immobilized on electrodeshave shown that much of the CYP immobilized on the electrode is in an inactive formand call into question the results from previous CYP biosensor studies162-164.

In addition, some CYP biosensor studies are missing crucial substrate turnover dataand instead rely on electrochemical experiments alone to indicate that the immobilizedCYP is capable of electrochemically- driven catalysis157.

31

33 | P a g e

Table 1-6 Summary of different electrode types and electrode modifications used to construct CYP electrodes and biosensors 165.

CYP enzyme Electrode material Electrode modification Result Reference

Adsorption to bare electrodes

CYP101 (P450CAM) Edge-Plane Pyrolytic

Graphite

None Reversible peaks for FeIII/FeII seen in anaerobic CV

Increased reduction peak area in presence of D-(þ )-

camphor

Kazlauskaite et al., 1996

CYP101 Mutant SCF-

K344C

Gold None Enhanced electroactivity of the SCF-K344C mutant Lo et al., 1999

CYP2E1 Gold None Reversible peaks for FeIII/FeII seen in anaerobic CV,

ks=5 s-1

Fantuzzi et al., 2004

CYP199A2 Basal-Plane Pyrolytic

Graphite

None Quasi-reversible peaks for FeIII/FeII seen in anaerobic

CV, ks=550 s-1

Fleming et al., 2007

CYP101 Glassy Carbon None Increased reduction peak area in presence of D- (þ )-

camphor, ks=0.016 s-1

Mhaske et al., 2010

Layer-by-Layer Adsorption

CYP101 Gold Multilayer films of PSS and PDDA

on an MPS SAM

Reversible peaks for FeIII/FeII seen in anaerobic CV

Electrochemically-driven styrene epoxidation

(turnover 9.3 h-1)

Lvov et al., 1998

CYP3A4 Gold MPS SAM followed by PDDA Reversible peaks for FeIII/FeII seen in anaerobic CV

Electrochemical detection of the 3A4 substrates

verapamil and midazolam

Joseph et al., 2003

CYP1A2 Carbon Cloth Alternate adsorption of PSS and 1A2 Reversible peaks for FeIII/FeII seen in anaerobic CV


(turnover 39 h-1)

Estavillo et al., 2003

CYP1A2 and CYP3A4

microsomes

Pyrolytic Graphite Multilayer films of PEI and PSS Reversible peaks for FeIII/FeII seen in anaerobic CV


Sultana et al., 2005

CYP1A2 and CYP2E1 Basal-Plane Pyrolytic

Graphite

Multilayer films of CYP and CPR/b5 Reversible reduction and oxidation peaks visible in

anaerobic CV

Krishnan et al., 2011

34 | P a g e

Evidence that electron transfer follows natural

catalytic pathway from CPR

Electrochemically-driven catalysis of NNK to HPB

by 2E1 electrode

Adsorption to Thin Films

CYP101 Glassy Carbon Pretreated sodium montmorillonite

clay colloid


Fast electron transfer (5 to 152 s-1 for scan rates of 0.4

to 12 V/s)

Lei et al.,2000

CYP2B4 Glassy Carbon Mixture of sodium montmorillonite

clay colloid, 2B4, and Tween 80


Amperometric detection of the 2B4 substrates

aminopyrine and benzphetamine

Shumyantseva etal., 2004

CYP101 Pyrolytic Graphite DDAB or DMPC Reversible peaks for FeIII/FeII seen in anaerobic CV

Relatively fast electron transfer with DDAB (ks=26 s-

1) and DMPC (ks=25 s-1)

Zhang etal.,1997

CYP2C9, 2C18, and

2C19

Edge-Plane Pyrolytic

Graphite

DDAB Reversible peaks for FeIII/FeII seen in anaerobic CV

for all three enzymes Weak signal-to-background

current and only 1–3% of enzyme is electroactive

Shukla etal.,2005

CYP2C9 Edge-Plane Pyrolytic

Graphite


Anodic shift in redox potential with 2C9 substrates

torsemide, warfarin, and tolbutamide and with CO

Johnson etal.,2005

CYP27B1 Edge-Plane Pyrolytic

Graphite


(ks=3.5 s-1)

Rhieu etal.,2009

No product formation observed during electrolysis

with 27B1 substrate 25(OH)D3

CYP3A4 fusion protein Glassy Carbon PDDA Electrochemically-driven catalysis with the 3A4

substrate erythromycin

Dodhia etal.,2008

CYP101 Glassy Carbon Covalent attachment to thin film of

pyrene maleimide

Increased reduction peak area in the presence of D-(þ

)-camphor

Mhaske etal.,2010

Screen-Printed Electrodes

35 | P a g e

Riboflavin-conjugated

CYP1A2, 2B4, and 11A1

Rhodium-graphite Mixture of CYP, BSA,

glutaraldehyde, sodium cholate, and

phospholipid

Electrochemically-driven catalysis with respective

1A2, 2B4, and 11A1 susbstrates


Amperometric detection of the substrates aminopyrine

(2B4) and cholesterol (11A1)

CYP1A2 Riboflavin-graphite Glycerol and agarose Amperometric detection of the 1A2 substrate

clozapine

Antonini etal.,2003

CYP11A1 Rhodium-graphite Gold nanoparticles Amperometric detection of the 11A1 substrate

cholesterol Enhanced electroactivity and sensitivity

with inclusion of gold nanoparticles


CYP2B4 Carbon Covalent attachment via EDC/NHS

to a COOH-functionalized electrode

Amperometric detection of the 2B4 substrate cocaine

in confiscated cocaine samples

Asturias-Arribas et al.,2011

CYP1A2, 2B4, and 51b1 Graphite Mixture of gold nanoparticles and

DDAB

Increased reduction peak area in the presence of

substrates for 2B4 (benzphetamine) and

51b1(lanosterol) Enhanced electroactivity with

inclusion of goldnanoparticles


CYP2B4 Graphite Mixture of gold nanoparticles and

DDAB

Electrochemically-driven catalysis with

benzphetamine exhibits similar uncoupling behavior

to catalysis with 2B4 in solution

Rudakov etal.,2008

CYP2B4, 3A4, 11A1,

51b1

Graphite Mixture of gold nanoparticles and

DDAB

Amperometric detection of multiple substrates and

inhibitors

Shumyantseva et al., 2011

Encapsulation in Polymers of Gels

CYP101 IndiumTinOxide Polypyrrole Electrochemically-driven catalysis with D-(þ)-

camphor

Sugihara et al.,1998

CYP102 (P450BM3)

Mutant

Platinum and Glassy Carbon Polypyrrole Compared chemical vs. Electrochemical

polymerization of polypyrrole for effect on enzyme

catalysis

Holtmann et al., 2009

CYP2B4 Gold Polypyrrole Amperometric detection of the 2B4 substrate

phenobarbitol

Alonso-Lomillo et al.,2008

CYP101 Glassy Carbon Methyltriethoxysilane sol-gel and

DDAB


in aqueous and organic media

Iwuoha et al.,2000

Amperometric detection of camphor and pyrene

36 | P a g e

CYP2B6 Glassy Carbon Mixture of chitosan and gold Enhanced electroactivity within clusion of chitosan

and goldnanoparticles

Liu etal.,2008

Electrochemically-driven catalys is of the 2B6

substrate bupropion

Covalent Attachment to Self-Assembled Monolayers on Gold

CYP2C9 Gold Amine coupling via EDC/NHS to a

mixed SAM of OT and MUA

Quasi-reversible peaksforFeIII/FeII seen in anaerobic

CV

Yang et al.,2009

Electrochemically-driven catalysis with the 2C9

substrate warfarin

CYP3A4 fusion protein Gold Amine coupling via EDC/NHS to a

mixed SAM of 6HT and 7MHA

Microfluidic cell made of the 3A4 electrode

amperometrically detected the 3A4 substrates

quinidine, nifedipine, alosetron, and ondansetron

Fantuzzi et al.,2010

CYP2C9n1, 2C9n2, and

2C9n3 CYP2D6n1,

2D6n2, and 2D6n17

Gold Amine coupling via EDC/NHS to a

mixed SAM of 6HT and

7MHA(2C9)Maleimide/thiol

coupling to maleimide-terminated

SAM (2D6)

Micro-machined eight-electrode array used to

amperometrically measure the KM and kcat values for

the 2C9 substrate warfarin and the 2D6 substrate

bufuralol

Panicco et al.,2011

CYPc17 Gold His-tag attachment via modified Ni-

NTA on na MPASAM in the

presence of DDAB

Redox peaks for FeIII/FeII seen in anaerobic CV Johnson and Martin, 2005

Enhanced electroactivity when using His-tag

attachment vs.amine-coupling

CYP2E1 Gold Maleimide/thiol coupling to

maleimide-terminated SAM

Redox peaks for FeIII/FeII seen in anaerobic CV Faster

(ks=10 s-1 ) electron transfer with maleimide/thiol

coupling compared to adsorption to thin films

Fantuzzi et al.,2004

CYP2E1 Single-Cysteine

Mutants

Gold DTME SAM Electrochemically-driven catalysis with the 2E1

substrate p-nitrophenol

Mak et al.,2010

CYP3A4 fusion protein Gold Maleimide/thiol coupling to

maleimide-terminated SAM

Electrochemically-driven catalysis with the 3A4

substrate erythromycin

Dodhia et al.,2008

37 | P a g e

Future Directions/Nanostructured Electrodes

CYP11A1 Rhodium-graphite MWCNTs on screen-printed

electrodes

Enhanced amperometric detection (improved

linearity) of cholesterol within clusion of MWCNTs

Carrara et al.,2008

CYP2B4, 2C9,3A4 Graphite or Gold MWCNTs on screen-printed

electrodes

Amperometric detection of benzphetamine (2B4),

naproxen (2C9), and cyclophosphamide

Enhanced limits of detection with inclusion of

MWCNTs

Cararra, et al.,2011

CYP3A4 Sputtered Gold Adsorption to thin film of napthalene

thiolate SAM

Enhanced electroactivity on sputtered gold electrodes

(ks=340 s-1)

Mie et al.,2010

CYP3A4 microsomes Microfabricated Gold

Nanodomes

Adsorption to thin film of napthalene

thiolate SAM

Enhanced electroactivity on nanostructured gold

electrodes

Ikegami et al.,2011

CYP3A4

Gold Amine coupling to ZnSe quantum

dots covalently coupled to na

MPASAM

Amperometric detection of the 3A4 substrate 17β-

estradiol

Ndangili et al.,2011

Anodic shift in the redox potential with inclusion of

quantum dots

1. INTRODUCTION 1.4. Heme proteins

1.4 Heme proteins

Both enzymes addressed in this thesis fall into the heme proteins category; CytochromeP450 (CYP) and Cytochrome c Nitrite Reductase (ccNiR).

Heme is perhaps the most ubiquitous cofactor found in nature and the most function-ally diverse. Proteins containing a heme group constitute a heterogeneous family whosefunctional impact in living organisms is exceedingly high, due to their involvement in alarge range of biological processes. By taking part in several metabolic processes withinliving creatures, heme proteins play a key role in the maintenance of life on earth 181 .Hemoproteins are involved in cell respiration (cytochromes), oxygen-binding and trans-port (hemoglobin and myoglobin), oxidative biotransformations (cytochrome P450 andperoxidases), and most recently discovered, as sensors in 2-component regulatory sys-tems (guanylate cyclase, FixL, and CooA).

The ability of hemoproteins to carry out extremely diverse reactions arises largelyfrom the protein environment in which the heme molecule resides and specifically thenature of the heme-ligands. Other factors that contribute to the reactivity of the hemeare intrinsic to the heme itself, including the substituents on the heme periphery and, insome cases, the covalent attachment of the heme to the protein182.

37

1. INTRODUCTION 1.4. Heme proteins

38

2Experimental

2.1 Part A- Cytochrome P450 1A2

The cytochrome P450 1A2 (protein 10 mg/mL; CYP1A2 122 pmol/mg) was providedby the Centro de Investigação em Genética Molecular Humana from IHMT. The proteinwas purified as previously described in Palma et al.183, and stored in TGE buffer (75 mMTris.HCl, 10 % (v/v) Glycerol and 25 mM EDTA, pH 7,5) at -80 C.

Caffeine was purchased from Sigma-Aldrich, the solutions were prepared in deion-ized water (18MΩ/cm) from Millipore MilliQ or Purelab Option (Elga) water purificationsystems.

2.1.1 Electrochemical measurements

A conventional three-electrode electrochemical cell was used, with an Ag/AgCl referenceelectrode, a Pt counter electrode and a PGE (φ=3mm) modified with the enzyme/sol-gelfilm, as working electrode. The electrochemical cell, containing 10mL of 0.5M PBS buffer,pH 7 as supporting electrolyte, was thoroughly purged with argon before and an ar-gon atmosphere was maintained inside the cell during the experiments. Measurementswere performed with a potentiostat Autolab PSTAT 12 (Eco-Chemie) monitored by thecontrol and data acquisition software NOVA 1.6. In cyclic voltammetry experiments, a50mV/s scan rate was used unless otherwise stated. All potentials were quoted againstthe Ag/AgCl reference electrode.

39

2. EXPERIMENTAL 2.1. Part A- Cytochrome P450 1A2

2.1.2 Mediated electrochemistry of cytochrome P450

Before modification the PG working electrodes (self-made with 3 mm diameter disksin glass tubes) were polished with alumina slurry (0.3 µm) from Buehler, thoroughlywashed with water and ethanol and ultra-sonicated in deionized water for 5 min. Finally,the electrodes were washed with water and dried with compressed air.

The mediators used, 1,1’-dimethyl-4,4’-bipyridinium dichloride (methyl viologen, MV)and 3,7-bis(Dimethylamino)-phenothiazin-5-ium chloride (methylene blue MB) were pur-chased from Sigma. All chemicals were of analytical grade.

Table 2.1: Mediators structure and formal reduction potential.

Mediator structure E°' (mV)vs NHE

Methylene

Blue184

-426

Methyl

Viologen130

-440

2.1.2.1 Mediator in solution

The electrode was prepared by depositing 7.5 µL of CYP1A2 on the PGE and drying atroom temperature for ca. 45 min. The assays were performed in the presence of 100 mMof mediator (either MB or MV) in the electrochemical cell.

2.1.2.2 Electropolymerized mediator

Electropolymerization of methylene blue was performed by cyclic voltammetry (11 scans);the CVs were traced at a scan rate of 100 mV/s in the potential window -0.6 to 1.2 V vsAg/AgCl , mediator concentration in solution was 100 mM.

After the mediator polymerization, a 7.5 µL drop of CYP1A2 was applied to the PGEand left to dry at room temperature for ca. 45 min.

2.1.3 Direct electrochemistry

2.1.3.1 Direct adsorption

The PG working electrodes were cleaned as described above, subsequently a 7.5 µL dropof CYP1A2 was applied on the electrode surface and left to dry at room temperature forca. 45 min.

40


2.1.3.2 Surfactant films

The surfactant films were usually prepared from aqueous solutions which were cast ontothe electrode surfaces. After solvent evaporation, a water-insoluble multi-bilayered filmstructure was formed (selfassembled). The structures of the surfactants used in the workdescribed in this section are shown in Figure 2.1.

Figure 2.1: Surfactant structures. Cationic head group: A) CTAB, B) DDAB; C) DDM(n-dodecyl-b-D-maltoside).

The protein (CYP1A2) was confined within different surfactant films (Dodecyldimethy-lammonium bromide (DDAB) and Cetyl Trimethyl Ammonium Bromide (CTAB) pur-chased from Sigma-Aldrich and n-Dodecyl β-D-maltoside (DDM) from CalBiochem R©),on the surface of basal-plane graphite electrodes (PGE) which were analyzed by cyclicvoltammetry. All chemicals were of analytical grade.

CYP1A2–surfactant solutions were prepared by mixing 1.0 mg/mL of enzyme with 2times the CMC of each surfactant (see Table 2.2). These mixtures were then incubated atroom temperature (22 ± 2 C) for 30 minutes. A 7.5 µL drop of mixture was then appliedto the PGE. The electrodes were dried at room temperature for ca. 45 min.

Table 2.2: Surfactant critical micelar concentrations.

41


2.1.3.3 Sol-gel matrices

Sodium Silicate (Figure 2.2A) (Sigma-Aldrich), Tetraethoxysilane (TEOS, 98%, Alfa Ae-sar) and Tetramethyl orthosilicate (TMOS 98%, Sigma-Aldrich) were used as the silicaprecursor. Poly(ethylene glycol) (PEG, MW 400 and 6000)(Figure 2.2B) were purchasedfrom Prolabo.

A B

Figure 2.2: A) structure of sodium silicate; B) structure of poly(ethylene glycol).

Phosphate buffer solutions (PBS) were prepared with Na2HPO4•2H2O (99.5%, Merck)and KH2PO4 (99.9%, Prolabo). Tris–HCl buffers were prepared using Tris(hydroxymethyl)aminomethane(Sigma, 99.8%) and HCl (36%, Prolabo). All solutions were prepared with high puritywater (18MΩ/cm) from a Purelab Option water purification system (Elga).

The PG working electrodes were cleaned as described above. For the sol-gel modifiedelectrodes the surfaces were first treated by abrasion with sandpaper followed by theregular procedure. The optimized hybrid sol-gel was prepared by mixing Sodium Silicate55 mM and PEG400 6% at pH 7. Protein films were applied onto the electrode surface byplacing 10 µL of a solution containing equal parts of CYP1A2 and sol-gel. The electrodeswere left to air dry for 1h.

Table 2.3: Compositions of sol-gel based matrices using TMOS and CTAB.

42

2. EXPERIMENTAL 2.2. Part B- Cytochrome c Nitrite Reductase

Table 2.4: Compositions of sol-gel based matrices using Sodium silicate and CTAB.

Table 2.5: Compositions of sol-gel based matrices using Sodium silicate and PEG400.

2.2 Part B- Cytochrome c Nitrite Reductase

The protein Cytochrome c Nitrite reductase from Desulfovibrio desulfuricans ATCC 27774was purified at Requimte, FCT-UNL, according to the procedure described by Almeidaet al.149 and stored in 0.1 M phosphate buffer pH7.6, at -20C. The specific activity was300 U/mg and the protein content 1.0 mg/mL.

2.2.1 Electrochemical measurements

conventional three electrode electrochemical cell was used, with an Ag/AgCl referenceelectrode (Radiometer), a Pt counter electrode (Radiometer) and a PG electrode (φ= 3mm)modified with the sol- gel/ccNiR film, as working electrode. The electrochemical cell,containing 10 mL of PBS pH 7.0 as supporting electrolyte, was thoroughly purged withargon before and an argon atmosphere was maintained inside the cell during the ex-periments. Measurements were carried out with a potentiostat Autolab PSTAT 12 (Eco-Chemie) monitored by the control and data acquisition software NOVA 1.6 (EcoChemie).Cyclic voltammetry experiments were performed with a 50 mV/s scan rate in a potentialwindow of -0.1 to -0.8 V.

2.2.2 Response to nitrite

To evaluate the biosensors response to nitrite, the electrochemical cell was successivelyspiked with standard solutions of sodium nitrite (0.01, 0.1 and 1 M). After each addi-tion, the cell was again purged with argon for ca. 30 s and the CV was recorded. Nitrite

43


catalytic currents were determined at the inversion potential (-0.8 V) unless stated other-wise. All potentials were quoted against the Ag/AgCl reference electrode (-197 mV vsNHE).

2.2.3 Hybrid sol-gel matrix

The sol-gel precursor Sodium Silicate was purchased from Sigma-Aldrich and the Poly(ethyleneglycol) (PEG, MW 400 and 6000) was acquired from Prolabo. Solutions were preparedwith deionized water (18 MΩcm) from a Millipore MilliQ water purification system. Allchemicals were analytical grade and were used without further purification.

The PGE working electrodes were cleaned as described in section 2.1.2). Protein filmswere prepared by depositing a 10 µL drop of the preparation, containing the enzymesolution (1.0 mg/mL) and the sol-gel (sodium silicate and PEG, either or kDa, were pre-pared according to the formulation described in Table 2.5 with a ratio of 44/5, on theelectrode surface. After 60 minutes, the electrode was washed with buffer and placed inthe electrochemical cell.

2.2.4 Macroporous Carbon Nanotubes

ingle-walled carbon nanotubes functionalized with carboxylic groups (CNT, >90%, 4–5nm 0.5-1.5 lm) were obtained from Sigma. Polystyrene beads (PS, φ ≈ 500 nm, 50mg/mL)were prepared by emulsion polymerization in water solution following a protocol fromthe literature M. Nagai et. al.185.

Glassy carbon plates used as working electrodes (GC, Sigradur R©, HTW Hochtemperatur-Werkstoffe, Germany) were polished before and after modification by wet emery paper4000 with Al2O3 powder (0.05 µm, Buehler).

Electrophoretic deposition of carbon nanotubes and polystyrene-beads was performedusing a cell consisting of stainless steel plate as cathode and glassy carbon plate as anode.The two electrodes were placed parallel to each other and separated by 6 mm Figure 2.3.The area of contact of each electrode with the dispersion was 1 cm2. The adequate voltagewas applied using a DC power supply.

Prior to enzyme coating, the GC plates working electrodes previously functionalizedwith carbon nanotubes (as described in section 2.1.2) were rinsed with deionized water.A drop of enzyme (7 µL) was then applied onto the modified electrodes which were leftdrying for 40 min under an argon flux.

The carbon nanotubes dispersion (0.1 mg/mL) was obtained by sonication in an ul-trasonic bath for 12 hours. The PS-beads were added under gentle stirring to the carbonnanotubes dispersion prior to the deposition in order to achieve a final concentration of0.025mg/mL . The two parallel electrodes were introduced in a 4mL aliquot of the CNTor CNT-PS dispersion and a constant voltage of 60 V was applied for the required time.The electrode dipping was kept constant in order to ensure the same area of contact with

44


the dispersion and therefore improve the reproducibility of the deposition. After depo-sition, the glassy carbon plate was carefully removed from the remaining dispersion anddried horizontally at room temperature. The electrodes prepared in such way were there-after after called GC-CNT or GC-CNT-PS. The template (PS-beads) removal was carriedout in an oven at 450 Cfor 1 hour with 15 C/min ramp resulting in electrodes calledGC-CNT-(macroporous).

Figure 2.3: Scheme of the experimental device used for the electrophoretic deposition ofcarbon nanotubes201.

A three electrode electrochemical cell was used Figure 2.4, with an Ag/AgCl referenceelectrode (Radiometer), a Pt counter electrode (Radiometer) and a modified glassy carbonplate as working electrode. The electrochemical cell containing 0.1 M KCl in 0.05 M Tris-HCl buffer pH 7.6 as supporting electrolyte, was thoroughly purged with argon beforeand during the experiments. Measurements were carried out with a potentiostat AutolabPSTAT 12 (EcoChemie) monitored by the control and data acquisition software NOVA 1.6(EcoChemie). Cyclic voltammetry experiments were performed with a 50 mV/s scan ratein the potential window -0.1 to -0.8 V.

To evaluate the biosensors response to nitrite, the electrochemical cell was succes-sively spiked with standard solutions of sodium nitrite (0.01, 0.1 and 1 M). After eachaddition, the cell was again purged with argon for ca. 30 s and the CV (cyclic voltammo-gram) was recorded. Nitrite catalytic currents were determined at the inversion potential(-0.8 V) unless stated otherwise. All potentials were quoted against the Ag/AgCl refer-ence electrode (-197 mV vs NHE).

The electrochemical experiments were performed according to section 2.2.1) exceptfor a few changes: namely, the electrochemical cell (Figure 2.3); the working electrode,

45


and a modified glassy carbon plate as working electrode.

Work electrode

GC Plate connected to

aluminum foil

Reference

electrode Ag/AgCl

Counter electrode

Platinum

Figure 2.4: Electrochemical cell.

2.2.5 Oxygen scavenger system

Glucose oxidase (Type II from Aspergillus niger 17.3 U.mg-1) and catalase (from bovineliver, 2-5kU mg−1) were purchased as lyophilized powders from Sigma and used as re-ceived. All other chemicals were of reagent grade. Deionized water (18 MΩcm) was usedin all experiments.

Prior to modification the PG electrodes were cleaned as previously described. Proteinfilms were prepared by depositing a 10 µL drop of the preparation, containing the en-zyme solution(s) and the sol-gel, on the electrode surface. After 60 minutes of air drying,the electrode was washed with buffer and placed in the electrochemical cell.

The enzymatic scavenger system constituted by glucose oxidase and catalase wastested in different ways: 1) with sol-gel immobilized ccNiR (4.5 U) and GOx (9.3 U/mL)and catalase (650 U/mL) in solution in the electrochemical cell; 2) sol-gel immobilizedccNiR (4.5 U) and catalase (41.7 U) with GOx (9.3 U/mL) in solution and.

46

3Results and discussion

3.1 Cytochrome P450 1A2

Aiming at the development of an electrochemical CYP1A2 based biosensor, P450 enzymewas used either in presence or absence of its natural redox partner (CYPOR complex).The different strategies were investigated using cyclic voltammetry. The experimentaldesign has taken into account the need to establish an effective electronic communicationbetween the enzyme and the electrode and, at the same time, preserve the biologicalactivity. It is important to note that to probe the catalytic activity of CYP1A2, caffeinewas selected from a plethora of substrates due to its low cost, availability and moderatewater solubility. In general, the co-substrate - molecular oxygen - was provided to themedium dissolved in the caffeine aqueous stock solutions or, alternatively, in deionizedwater. Unless stated otherwise the experiments were carried out in a purged electrolytesolution in order to remove oxygen which exhibits a broad waved that might mask thenon-catalytic signals of P450.

3.1.1 Cytochrome P4501A2 in the absence of CPR

The following section describes several strategies tested with the purpose of establishingthe electronic communication between the cytochrome and the electrode. At an initialstage, a mediated approach was used that later proved itself inadequate to our goals.This approach was discarded in favor of direct electrochemistry.

47

3. RESULTS AND DISCUSSION 3.1. Cytochrome P450 1A2

3.1.1.1 Mediated electrochemistry

Selection of a suitable mediator Three mediators were selected and tested as elec-tron shuttles, namely methyl viologen, methylene blue and methylene green (either inmonomeric or polymeric forms). These substances have sufficient negative potentialscompared to the redox potentials of the active sites of P450 (-0.4V ± 0.1V) and mightreduce the oxidized form (PFeIII) of the enzyme. In a first approach CVs were recordedwith the CYP immobilized on the PG electrode in the presence of dissolved mediators.

Following the addition of a caffeine stock solution, catalytic reduction currents couldclearly be observed (Ic increases while Ia decreases); to illustrate this behavior Figure 3.2and Figure 3.3 show the CV of methyl viologen and methylene blue in the presence ofan electrode with membrane entrapped-CYP. This means that the reduced forms of eachmediator were able to deliver electrons to a CYP which in turn, passes the electrons tothe substrate (MV Figure 3.2 and MB Figure 3.3). However, control experiments donein the absence of caffeine (additions of non-purged water aliquots) produced a similarbehavior. It became clear that the catalytic response was due an increase in the oxygenconcentration (water dissolved) and not the caffeine’s (Figure 3.1). On the other hand,control experiments performed in the absence of protein did not show significant currentvariations. Apparently, despite CYP being able to reduce molecular oxygen, the organicsubstrate caffeine is not participating in the catalytic reaction.

Figure 3.1: Schematic representations of the working principles of a CYP Bioelectrodewith mediated transduction.

Because at this stage the focus was on the catalytic response of CYP to caffeine only,this approach was abandoned.

48


-1.0 -0.9 -0.8 -0.7 -0.6 -0.5 -0.4

-6.0

-5.0

-4.0

-3.0

-2.0

-1.0

0.0

1.0

2.0

0 10 20 30 40 50

-1.00

-2.00

-3.00

-4.00

-5.00

Cu

rre

nt (

A)

E versus Ag/AgCl (V)

Ip (

mA

)

[caffeine] (mM)

Caffeine

solution

A B

Figure 3.2: A) Cyclic voltammograms of methyl viologen in the presence of CYP mem-brane entraped on a PG electrode, buffer solution 0.1 M MV, 0.1 M KCl and tris-HClbuffer 50 mM pH 7.6. in the presence of varying caffeine concentrations (0-42mM).Scanrate: 50mV s−1.B Icat variation with nitrite concentration.

A B

-0.5 -0.4 -0.3 -0.2 -0.1 0.0

-14

-12

-10

-8

-6

-4

-2

0

2

4

6

8

10

0.00 0.01 0.02 0.03 0.04 0.05

-0.90

-1.00

-1.10

-1.20

-1.30

-1.40

-1.50

Cu

rre

nt (

A)


Ip (A

)

[caffeine] (mM)

Caffeine

solution

Figure 3.3: 3 A) Cyclic voltammograms of methylene blue in the presence of CYP on aPG electrode, buffer solution 0.1 M MV, 0.1 M KCl and tris-HCl buffer 50 mM pH 7.6. inthe presence of varying caffeine concentrations (0–42mM).Scan rate: 50mV s−1.B . B) Icatvariation with nitrite concentration.

49


3.1.1.2 Direct electrochemistry of cytochrome P450

Direct electrochemistry of cytochrome P450 in surfactant films Because the electro-chemically mediated experiments did not give the expected full catalytic response, it wasdecided to use a DET approach. In fact, direct electrochemical reduction has the potentialto provide P450s with reducing equivalents, thereby eliminating the native ET machinery.

The first attempt at direct electrochemistry was carried out in the absence of surfac-tants or other stabilizing components. The successive voltammograms depicted in Figure3.4A show the change in the non-catalytic signals of cytochrome P450 1A2 with time.

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-0.40

-0.35

-0.30

-0.25

-0.20

-0.15

-0.10

-0.05

0.00

0.05

0.10

0.15

0.20

Curr

ent

(A

)


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-0.40

-0.35

-0.30

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-0.20

-0.15

-0.10

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0.00

0.05

0.10

0.15

0.20

Curr

ent

(A

)


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-0.40

-0.35

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-0.20

-0.15

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0.00

0.05

0.10

0.15

0.20

Curr

ent

(A

)


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-0.40

-0.35

-0.30

-0.25

-0.20

-0.15

-0.10

-0.05

0.00

0.05

0.10

0.15

0.20

Curr

ent

(A

)


A B

DC

Figure 3.4: Consecutive cyclic voltammograms of CYP1A2–surfactant films casted on PGelectrodes in 0.1 M KCl and tris-HCl 50 mM pH 7.6; scan rate 50 mV/s. A) Controlelectrode with CYP1A2 only, B) DDM C) DDAB, D) CTAB.

Although a small and broad redox wave could be observed in the range -0.3;-0.5 V thesignal promptly disappeared; this signal is similar to the ones observed in electrochemi-cal experiments using other cytochromes P45075,163,186,187, leading to the conclusion thatthis non-catalytic signal is correlated with the reduction and reoxidation of P450FeIII/FeII.This signal was not systematically observed and, when observed, was of very small in-tensity, probably due to of the small amount of protein that was able to reach the re-quired orientation to keep the conformation for direct electron transfer reaction and/orto a small ET rate.

In order to improve the protein stabilization and subsequent direct electron transferit was decided upon the use of surfactants. This CYP is a membrane-bound protein forwhich a more hydrophobic environment could be beneficial. Surfactant assemblies oncarbon electrodes have been shown to facilitate direct ET with various P450s74,188,189. The

50


protein was therefore incorporated into surfactant layers deposited on the electrode.

All the surfactants tested were used in a concentration corresponding to 2xCMC toensure that micelles could be formed on the electrode and, in this way, equivalently com-pare the effects of the surfactants on the enzyme’s electrochemical response.

Cyclic voltammograms of CYP1A2–surfactant films casted on PG electrodes are shownin Figure 3.4. The peak potential was never shifted, no matter the surfactant type. Thevoltammograms depicted in Figure 3.4 show a consistency in the electrochemical behav-ior of the protein in different surfactant films. It is easier to observe in Figure 3.4 B and D,that the peak potential is in the -0.350 to -0.400V range, and that it shifts slightly to morenegative potentials over time.

The reported studies on heme proteins, like hemoglobin and myoglobin, consideredthat the DDAB surfactant films could favor a specific orientation of the proteins embed-ded in them, due to interaction with the cationic head group or the hydrocarbon chainsof DDAB63,68. However, in this study DDAB did not provide the best results. On thecontrary, DDM and CTAB enabled a better electrochemical response. CTAB in particu-lar, a cationic surfactant such as DDAB, greatly enhanced the DET of CYP with the PGelectrode, therefore, it was selected for further studies.

The effect of the scan rate is shown in Figure 3.5. The cathodic peak currents in-creased linearly with the scan rate up to 1 V/s which is consistent with a diffusionlesselectrochemistry (Figure 3.5); only CVs with a scan rate up to 0.5 V are shown). The ab-sence of the symmetrical counterpart of the peak located at -375 mV indicates that theelectrode reaction is nonreversible.

0 100 200 300 400 500

0.00

-0.05

-0.10

-0.15

-0.20

-0.25

Ip (A

)

scan rate (mV/s)

Figure 3.5: Variation of the cathodic peak currents on the potential with the scan rate(5-500 mV/s) (y=-2.48*10-8-4.48*10−10x; r2=0.992)

51


The catalytic activity of the CYP1A2-CTAB films towards caffeine was investigated;substrate solutions were deoxygenated before addition to the electrochemical cell. Onceagain, no relevant changes could be detected in the voltammograms, i.e., there was nocatalytic activity (results not shown).

52


Stabilization of membrane-bound P450 protein for bioelectrochemistry

Sensor optimization The optimum membrane composition was selected from agroup of sol-gel formulations prepared with different precursors, water: monomer molarratios and enzyme quantities. The bioelectrodes were characterized by cyclic voltamme-try.

The first efforts were based on formulations containing CTAB, due to the results ob-tained in the previous section. The formulations based on the sol-gel precursors TEOSand TMOS had a tendency to precipitate, regardless the solvent/precursors/surfactantratios tested (Table 2.3). The sol-gel films had a thin and glassy appearance, but a hightendency to fracture during the drying process, occasionally loosing small pieces of glass.For these reasons these precursors were discarded and it was decided to test a differenttype of precursor, sodium silicate.

The sol-gel matrices prepared by the combination of sodium silicate and CTAB (Ta-ble 2.4) provided fairly more positive results (20 mM sodium silicate, 5 mM CTAB). Still,the film was thin and brittle. Although it was possible to obtain an electrochemical re-sponse, it was lost once the film cracked. Besides PEG being known to improve sol-gelconsistency, it has also been reported as a stabilizing agent for P450 in electrochemical ex-periments at high temperature190. Thus, it was tested here to immobilize CYP1A2 (Figure3.6C). However, no signal was observed in these conditions. The immobilization in sili-cate was also considered (Figure 3.6B), leading to a slight improvement in the measuredelectrochemical signal.

Next, sol-gel/enzyme films were prepared with the combination sodium silicate andpolyetilene glycol (Table 2.5). With the optimal formulation: 55 mM sodium silicate and6.25% PEG400 it was possible to obtain a hybrid film stable enough to continue our work.These results lead to the assumption that the use of sodium silicate prevents the introduc-tion of alcohol in the starting sol, while PEG improves the stabilization of the membraneprotein. This hybrid sol-gel formulation has not been previously reported and its reactionmechanism is not yet been established.

As previously stated, only the combination of PEG and silicate led to well definedredox peak at -0.335 V versus Ag/AgCl. The redox signal decreased rapidly by a factorof three during the first minutes of placing the electrode in electrolyte solution (Figure3.7). Most likely, the weakly immobilized protein molecules are leaching out from thefilm and only the stably encapsulated CYP remains attached in the hybrid sol-gel film forlong time experiments. This set of results clearly show that neither PEG nor silicate areable to immobilize CYP1A2 in an electrochemically active form and only the synergeticeffect from their combination allows the redox activity of the membrane-bound protein.

53


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-0.25

-0.20

-0.15

-0.10

-0.05

0.00

0.05

0.10

0.15

0.20

Curr

ent

(A

)


-0.8 -0.6 -0.4 -0.2 0.0 0.2

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ent

(A

)


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ent

(A

)


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-0.10

-0.05

0.00

0.05

0.10

0.15

0.20

Curr

ent

(A

)


A B

DC

Figure 3.6: Cyclic voltammograms of CYP1A2 casted on PG electrodes, in 0.1 M KCland tris-HCl 50 mM pH 7.6 purged electrolyte; scan rate 50 mV/s, in the presence of Aonly CYP; B CYP/Sodium Silicate; C CYP/Peg400; D CYP/Sol-gel(sodium silicate andPEG400).

Characterization of the non-catalytic signals The bioelectrodes, prepared with thehybrid sol-gel formulation described in the previous section were, characterized by cyclicvoltammetry. A non-catalytic current-potential curve was always observed in purgedelectrolyte without caffeine (Figure 3.7); the peak potential is coherent with the one ob-tained in the presence of surfactants (previous section) as well as the potentials describedin the literature for the heme iron reduction191. However, according to the literature,the CYP1A2 electrochemical signal in the absence of oxygen should depict the reversibleelectrochemical conversion of PFeIII/PFeII indicating that perhaps the vestigial oxygenconcentration present in the sol-gel film/electrolyte is causing the anodic signal to disap-pear. This phenomena will be further discussed in the next section.

54


-0.8 -0.6 -0.4 -0.2 0.0 0.2 0.4

-0.4

-0.3

-0.2

-0.1

0.0

0.1

Cu

rre

nt

(A

)


Figure 3.7: Consecutive cyclic voltammograms of PG/CYP1A2–sol-gel film electrode in0.1 M KCl and 50 mM tris-HCl buffer pH 7.6 purged with argon. scan rate 50 mV/s.

The electrochemistry of sol-gel encapsulated CYP1A2 was studied by changing sys-tematically the potential scan rate from 5 to 1000 mV s-1 as reported in Figure 3.8. Onlyone peak could be observed, and its current intensity increased linearly with scan rate’ssquare root which is consistent with a diffusion controlled electrochemistry (Figure 3.8).The enzyme is thought to be immobilized in a matrix that does not prevent diffusionsince no deviation from linearity is observed between peak current and square root ofthe scan rate up to 1 V/s (Figure 3.8 only shows up to 0.50 V/s).

55


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-0.5

-0.4

-0.3

-0.2

-0.1

0.0

0.1

0.2

0.3

Cu

rre

nt

(A

)


0 5 10

-0.06

-0.08

-0.10

-0.12

-0.14

-0.16

-0.18

-0.20

-0.22

Ip (A

)

(scan rate)^(1/2) (mV/s)

A B

Figure 3.8: A) Cyclic voltammograms of PG/CYP1A2–sol-gel film electrode in 0.1 M KCland 50 mM tris-HCl buffer pH 7.6 purged electrolyte at different scan rates; from inside tooutside: 0.005, 0.01, 0.02, 0.05, 0.1, 0.2, 0.25 V.s−1, respectively. B) Variation of the cathodicpeak currents on the potential scan rate (y=-0.03-0.017x; r2=0.98).

Response to oxygen and caffeine The chemistry of oxygen activation catalyzed bythe P-450s has always been a topic of great controversy. The heme iron of P450 is coorde-nated by a cysteine as the fifth ligand. The sixth position is left vacant for oxygen bindingduring catalysis - but may be occupied by a water molecule in the resting state. The in-fluence of dissolved oxygen on the electrochemical response of CYP1A2 immobilized insol-gel (PEG400 and sodium silicate), observed in Figure 3.9 reveals that when the elec-trode is exposed to a high percentage of oxygen the cathodic signal, however intense inthe first scan, quickly disappears. The electrochemical response measured for the firstcyclic voltammogram showed a well-defined peak with a current intensity higher than 8µA and maximum peak potential at -0.355 V vs Ag/AgCl. Still, continuous cycling at 50mV s-1 leads to a continuous signal degradation and a peak shift to lower potentials. Ac-cording to Rusling191, after electrochemical reduction of P450-FeIII in aerobic solution,reduced form PFeII is combined with dioxygen to form PFeII -O2 complexes and thiscomplex might be further reduced electrochemically, yielding hydrogen peroxide and re-generating the PFeII heme192,193. The peroxide generated in this catalytic reduction ofoxygen reacts with cyt P450FeIII to form +•(P450-FeIV=O), which should be reduced atthe electrode to regenerate cyt P450-FeIII (Figure 3.10), the lectroactive species whose re-duction is being monitored. Though, this regeneration does not appear to occur, whichleads to the possible conclusion that the cycle described in the scheme is blocked in oneof the steps. In such a case there are three hypotheses:

56


i) The window of potential in which the reaction was analyzed is not negative enoughto reduce the +•(P450-FeIV =O), in other words, there is a lack of a driving force ableto reduce this form of P450 making it impossible to regenerate the PFeIII (step 6blocked);

ii) The PFeIII heme degradation due to hydrogen peroxide accumulation (c.f. 3.2.3)(step 5 blocked);

iii) The CYP’s conformational change necessary for P450-FeII -O2 conversion in hydro-gen peroxide and P450-FeII does not occur (step 3 blocked).

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-3.0

-2.5

-2.0

-1.5

-1.0

-0.5

0.0

0.5

Cu

rre

nt (

A)


Time

Figure 3.9: Consecutive cyclic voltammograms of PG/CYP1A2–sol-gel film electrode in0.1 M KCl and 50 mM tris-HCl buffer pH 7.6 aerobic conditions. scan rate 50 mV/s.

57


Figure 3.10: Pathways for Biocatalytic Activation of Cyt P450s by Peroxides, OxygenReduction Formed Peroxide.by P450s on Electrodes.

The results obtained upon the addition of purged caffeine aliquots turned out to beequivalent to the ones acquired when using redox mediators or DET with surfactants,meaning no catalytic response could be observed (Figure 3.11). This signal representsstep 1 (Figure 3.10) in the presence of small amounts of oxygen (vestigial). If the condi-tions were completely anaerobic the signal should be reversible. However, in the pres-ence of oxygen the equilibrium is shifted in the direction of PFeIIO2 formation, turningthe reaction (step 1) irreversible.

58


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-0.3

-0.2

-0.1

0.0

0.1

0.2

Cu

rre

nt

(A

)


Figure 3.11: Cyclic voltammograms of CYP1A2–sol-gel film casted on PG electrode,buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6 purged with argon. In the presenceof varying caffeine concentrations in the absence of oxygen. Scan rate: 50mV s−1.

Figure 3.12 reports the electrochemical response of CYP1A2 immobilized in the sol-gel-PEG hybrid film to an increasing amount of non-deareated caffeine. The experimentwas initiated in an oxygen free solution (purged with Argon). Then, small volumes of anon-purged caffeine solution were introduced in the cell, leading to the gradual increasein the cathodic peak intensity and a decrease in the anodic currents. It important tonote that the data was plotted against the volume of caffeine added, in order to have acomparison term with the experiments where water additions were made.

59


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-2.60

-2.40

-2.20

-2.00

-1.80

-1.60

-1.40

-1.20

-1.00

-0.80

-0.60

-0.40

-0.20

0.00

0.20

0.40

-200 0 200 400 600 800 1000 1200 1400 1600

0.00

-0.50

-1.00

-1.50

-2.00

Cu

rre

nt (A

)

Potential applied (V)

Ip (A

)

Volume added (L)

Caffeine

Solution

Figure 3.12: Cyclic voltammograms of CYP1A2–sol-gel film casted on PG electrode,buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6 purged with argon. In the presenceof varying non-purged caffeine volumes (stock solution 60 mM). Scan rate: 50mV/s.

Conversely, control experiments carried out adding equivalent volumes of deareatedwater, provided the same results revealing that the responses observed was due to anincrease in oxygen catalysis and not to caffeine’s (Figure 3.13). The following experimentwas initiated in an oxygen free electrolyte solution. When small non-purged water vol-umes were added to the solutions bathing CYP1A2/sol-gel/PG electrodes, the PFeIII

reduction peak increased greatly (Figure 3.13). This is the classic voltammetric signaturefor electrochemical catalysis. Here, it the reflects reaction of PFeII with dioxygen (step 2of Figure 3.10) followed by reduction of the PFeII -O2 complex to give H2O2 , regenerat-ing PFeII to continue the reaction cycle (step 3 of Figure 3.10).

60


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-2.8

-2.6

-2.4

-2.2

-2.0

-1.8

-1.6

-1.4

-1.2

-1.0

-0.8

-0.6

-0.4

-0.2

0.0

0.2

0.4

0 500 1000 1500 2000 2500 3000 3500

0.00

-0.50

-1.00

-1.50

-2.00

-2.50

Cu

rre

nt (

A)

E versus Ag/AgCl (V)Ip

(A

)Volume added (L)

Deionized

water

Figure 3.13: Cyclic voltammograms of CYP1A2–sol-gel film casted on PG electrode,buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6 purged with argon. Consecutiveadditions of non-purged water. Scan rate: 20mV s−1.

3.1.1.3 Catalase

As previously mentioned, hydrogen peroxide formation might be one of the reasons why,in the presence of high concentrations of oxygen, the catalytic peak, corresponding step1 (Figure 3.10), disappears after a few scans (Figure 3.9). In an attempt to avoid hemedegradation from hydrogen peroxide exposure194 several experiments were carried outin the presence of catalase. This enzyme is a commonly found in nearly all living organ-isms exposed to oxygen. A number of recent studies indicate that catalase is the primaryenzyme responsible for protecting the cells from hydrogen peroxide by catalyzing thefast decomposition of hydrogen peroxide to water and oxygen Equation 5.

H2O2 → H2O2 +O2 (3.1)

Table 3.1 shows the experiments performed to diminishing the peroxide’s impact inthe P450’s electrochemistry. It can be seen that the catalase addition to the cell doesnot prevent the decrease in the signal’s intensity, regardless some assumptions can bemade concerning the profile of the cyclic voltammograms. The twisted reverse tracephenomenon tends to increase in the presence of catalase at lower scan rates, accordingto what Limoges et al. reported194 this is characteristic of a chemical inactivation–redoxreactivation mechanism. Additionally, when looking at the negative sweep and consider-ing the results obtained by Butt et al.195 when studying Nitrite reductase in the presence

61


of inhibitors, is possible to infer that the combination of the magnetic stirring and thelower scan rate promotes hydrogen peroxide removal of the electrode surface.

62

63 | P a g e

Table 3-1 Compilation of several electrochemical experiments with sol -gel/CYP1A2 in anaerobic conditions.

50 mV/s without stirring 50 mV/s stirring 10 mV/s stirring

Without

catalase

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-10

-8

-6

-4

-2

0

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-10

-8

-6

-4

-2

0

Cu

rre

nt (

A)


Curr

ent

(A

)


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-8

-6

-4

-2

0

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-8

-6

-4

-2

0

Cu

rre

nt (

A)


Curr

ent

(A

)


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-5

-4

-3

-2

-1

0

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-5

-4

-3

-2

-1

0

Cu

rre

nt (

A)


Cu

rre

nt (

A)


With

catalase

1mg/mL

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-8

-7

-6

-5

-4

-3

-2

-1

0

1

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-8

-7

-6

-5

-4

-3

-2

-1

0

1

Cu

rre

nt (

A)


Curr

en

t (

A)


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-10

-8

-6

-4

-2

0

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-10

-8

-6

-4

-2

0

Cu

rre

nt (

A)


Cu

rre

nt (

A)


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-7

-6

-5

-4

-3

-2

-1

0

1

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-7

-6

-5

-4

-3

-2

-1

0

1

Cu

rre

nt (

A)


Cu

rre

nt (

A)



3.1.2 CYPOR complex

In order to better understand the results obtained with CYP1A2 namely, the lack of cat-alytic activity towards caffeine, similar experiments were performed with the CYPORcomplex. Considering that in vivo, as previously discussed in section 1.3.1), the CPRtransfers the electrons form NADPH to the cytochrome, it would be expected that, inthe presence of the cytochrome P450 reductase there could be catalytic activity towardscaffeine. For the immobilization of the CYPOR complex the same sol-gel was used as de-scribed in section 3.1.1.2)b)). The CVs revealed a cathodic peak at -0.2 V and a small an-odic wave at -0.33 V. As observed in Figure 3.14 the cathodic peak currents vary linearlywith the scan rate in a 5-500 mV/s range. This behavior indicates that the electrochemicalreduction of CYPOR is not controlled by diffusion inside the sol-gel film. Although thecyclic voltammograms depicted in Figure 3.14 do not fulfil the reversibility criteria, weare now able to observe an anodic peak, unlike what happened in the absence of the CPR.

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-0.40

-0.35

-0.30

-0.25

-0.20

-0.15

-0.10

-0.05

0.00

0.05

0.10

0.15

0.20

0.25

0.30

0 20 40 60 80 100

0.00

-0.02

-0.04

-0.06

-0.08

-0.10

Cu

rre

nt

(A

)


Ip (A

)

scan rate (mV/s)

Figure 3.14: A) Cyclic voltammograms of PG/CYPOR–sol-gel film electrode in 0.1 M KCland 50 mM tris-HCl buffer pH 7.6 in a purged electrolyte at different scan rates; frominside to outside: 0.005, 0.01, 0.02, 0.05, 0.1, 0.2, 0.25 V.s-1, respectively. B) Variation of thecathodic peak currents on the potential scan rate (y=1.85x10-10−8.39x10−10x; r2=0.997).

In aerobic conditions, the CYP bioelectrode’s CVs reveal a sharp catalytic peak. Heme–iron containing enzymes such as P450 have, not only, the ability to incorporate an oxy-gen atom from O2 into organic substrates, but also to utilize H2O2 and other peroxidesin the one-electron oxidation of the different cosubstrates (peroxidase activity)196. Thisfact should account for the heme protection from degradation while exposed to hydro-gen peroxide, but as it was discussed in the previous sections, this does not occur (Figure3.15B). Nonetheless, in the experiments containing CPR (Figure 3.15A) it is possible to

64


observe that the catalytic activity does not disappear. In fact, although a decrease of cur-rent intensity occurs in the first 10 cycles, at that point the signal stabilizes still displayinga “strong” catalytic activity, leading to the supposition that the CPR is somehow relatedto the peroxidase activity of the cytochrome P450.

-0.8 -0.6 -0.4 -0.2 0.0 0.2

-4.0

-3.0

-2.0

-1.0

0.0

Cu

rre

nt (

A)


-0.8 -0.6 -0.4 -0.2 0.0 0.2


a a

b b

A B

Figure 3.15: Cyclic voltammograms of films casted on PG electrode in aerobic condi-tions, buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6. Scan rate: 50mV s−1.of A)CYPOR–sol-gel B) CYP1A2–sol-gel film. a) 1st scan b) 20th scan.

The difference observed helps sustain the hypothesis postulated in the previous sec-tion, stating that a change in the CYP’s conformation is required in order to perform theP450-FeII -O2 conversion in hydrogen peroxide and P450-FeII (step 3, Figure 3.7); there-fore CPR is an essential element for the structural rearrangement. Upon the additionof small volumes of non-purged water to electrochemical cell an increase in the CYP’scatalytic response is observed (Figure 3.16).

65


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-2.8

-2.6

-2.4

-2.2

-2.0

-1.8

-1.6

-1.4

-1.2

-1.0

-0.8

-0.6

-0.4

-0.2

0.0

0.2

0.4

Cu

rre

nt (

A)


O2

Figure 3.16: The effect of O2 binding on the electrochemistry of the CYPOR/sol-gel castedon a PG electrode, purged buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6. Con-secutive additions of non-purged water. Scan rate: 50mV s−1.

Interestingly, unlike what was observed in the previous experiments carried out, inthe absence of CPR, when caffeine was added to the electrochemical cell, already con-taining a non-saturating oxygen concentration, a decrease in the catalytic current wasobserved (Figure 3.17). A similar catalytic current decreasing phenomenon has been de-scribed in the literature with glucose oxidase. Liu et al.197 for instance reported a de-crease in the catalytic response upon glucose addition. In the work reported, GOD(FADH2),electrocatalyzes the reduction of dissolved oxygen according to following equations.

GOD(FAD) + 2e− + 2H+ ↔ GOD(FADH2) (3.2)

GOD(FADH2) +O2 ↔ GOD(FAD) +H2O2 (3.3)

In the presence of glucose the electrocatalytic reaction is restrained due to the enzyme-catalyzed reaction between the oxidized form of GOD, GOD(FAD), and glucose, whichthen results in a decrease of electrocatalytic response (Equation 3.4).

Glucose+GOD(FAD)→ gluconolactone+GOD(FADH2) (3.4)

66


-0.8 -0.6 -0.4 -0.2 0.0 0.2

-0.14

-0.12

-0.10

-0.08

-0.06

-0.04

-0.02

0.00

0.02

0.04

0.06

0.08

Cu

rre

nt (

A)


Figure 3.17: Cyclic voltammograms of CYPOR/sol-gel films casted on PG electrode inanaerobic conditions, buffer solution, 0.1 M KCl and tris-HCl 50 mM pH 7.6 purged withargon.scan rate, 50 mV/s.(black-line) without substrates (red-line) oxygen addition (blueand green lines) caffeine additions increased concentration.

67

3. RESULTS AND DISCUSSION 3.2. Cytochrome c nitrite reductase

3.2 Cytochrome c nitrite reductase

n recent years, several ccNiR (D. desulfuricans ATCC 27774) based electrochemical biosen-sors have been proposed125. Herein, this well-established biosensor system was used as amodel to evaluate new immobilization platforms and operation methods. Unless statedotherwise, the experiments were carried out in a purged electrolyte solution in order toremove oxygen, which exhibits a broad waved that might mask the (non-) catalytic sig-nals of ccNiR. In this section, several approaches will be presented, whose purposes wereto improve the electronic communication and diminish the effect of interferences in theccNiR based sensor. The first strategy focused on the optimization of the use of the solgel formulation developed in section 3.1.1.2) using ccNiR as a model enzyme. In a secondapproach, the electrophoretic deposition of macroporous assemblies of single-walled car-bon nanotubes (SWCNTs) was used with the goal of improving the sensitivity of ccNiRbased nitrite biosensors. Lastly, an oxygen scavenger system was explored with the aimof avoiding the mandatory oxygen removal when working with reductase enzymes.

3.2.1 Hybrid sol-gel matrix

Like CYP1A2, ccNiR is a heme containing protein associated to the periplasmic mem-brane; for this reason, the matrix previously used for CYP immobilization was tested.This time, the sol-gel film was optimized by varying the polymer’s size.

3.2.1.1 Non catalytic signals

As shown in Figure 3.18, the CVs of the PEG400 based films have higher cathodic peakcurrents, which can indicate that this matrix is more suitable for the detection of theelectrochemical signals of ccNiR. The non-turnover currents should reflect the amount ofimmobilized enzyme molecules and their ability to interact with the electrode. The CVsof the PEG400 based films produce a higher cathodic peak currents, which can indicatethat this matrix is more suitable for the direct ET with ccNiR. Though, the results weresomewhat surprising because the higher polymer size of the PEG6000, is was expected toprovide larger pore sized films which constitute a more appropriate environment for theincorporation of the large molecular weight aggregates of ccNiR (as purified this enzymeis normally found is aggregates of over 890 kDa)149. In effect, other sol-gel formulationstested in the past124, using diferent monomer precursors indicated that those providingthe largest porosity delivered the best results. Such broad envelope comprises all hemesbelonging to both NrfA and NrfH subunits (14 hemes, per NrfHA trimer) with reductionpotentials ranging +150mV to 400mV vs NHE, as previously stipulated by protein filmvoltammetry, using an enzyme layer directly adsorbed on pyrolitic graphite electrodesurface. Since the potential of the cathodic peak was more negative when ccNiR wasentrapped in the PEG400 based formulations, perhaps in this case, the electrochemicalreaction beneath is related with the reduction of the more negative hemes. This might be

68


a consequence of different molecular reduction within this sol- gel matrix. Future surfacecharacterization of these two enzyme films should indicate if the protein is effectivelyencapsulated in both of them. If ccNiR is embedded only in the PEG6000 sol-gel, then thedirect electrode contact on the PG surface may be somewhat impaired, and in contrast,with the PEG400 sol-gel but heterogeneously deposited on the electrode surface.

-0.9 -0.8 -0.7 -0.6 -0.5 -0.4 -0.3 -0.2 -0.1 0.0

-0.12

-0.10

-0.08

-0.06

-0.04

-0.02

0.00

0.02

0.04

0.06

Curr

ent (

A)


-0.9 -0.8 -0.7 -0.6 -0.5 -0.4 -0.3 -0.2 -0.1 0.0

-0.12

-0.10

-0.08

-0.06

-0.04

-0.02

0.00

0.02

0.04

0.06

Curr

ent (

A)


A B

Figure 3.18: Cyclic voltammograms of ccNiR immobilized on sol -gel films, buffer solu-tion, 0.1 M KCl and tris-HCl 50 mM pH 7.6 purged with argon.scan rate, 20 mV/s. A)ccNiR entrapped in a sodium silicate and PEG400 film B) ccNiR entrapped in a sodiumsilicate and PEG6000 film.

69


3.2.1.2 Response to nitrite

To assess the dependence of the catalytic current on the concentration of the substrate,cyclic voltammograms of the PG/ccNiR-PEG modified electrodes were registered at var-ious nitrite concentrations. The correspondent cyclic voltammograms for a PG/ccNiR-PEG6000 electrode are presented in Figure 3.19; they show increased cathodic peak cur-rents correlated with the substrate amount. This reflects the electroenzymatic reductionof nitrite to ammonium and hence, a direct electrical connection of ccNiR to the electrodesurface.

-0.9 -0.8 -0.7 -0.6 -0.5 -0.4 -0.3 -0.2 -0.1 0.0

-3.00

-2.80

-2.60

-2.40

-2.20

-2.00

-1.80

-1.60

-1.40

-1.20

-1.00

-0.80

-0.60

-0.40

-0.20

0.00

0.20

Cu

rre

nt (

A)


Nitrite

0µM

500µM

Figure 3.19: Electrochemical response of PG/ccNiR–sol-gel film(PEG6000) electrode tovarying nitrite concentrations (0-500µM) in 0.1 M KCl and 50 mM tris-HCl buffer pH 7.6purged with argon. Scan rate, 20 mV/s.

Figure 3.20 shows the relation between the catalytic currents and nitrite concentrationfor ccNiR/PEG400 and ccNiR/PEG6000 electrodes. Apparently, the nitrite reducing ac-tivity follows a typical Michaelis-Menten behavior, i.e. a steady increase of the catalyticcurrents is observed, with a linear range extending up to 100 µM of nitrite and reachinga plateau at higher concentrations.

70


0 100 200 300 400 500

0.00

-0.50

-1.00

-1.50

-2.00

-2.50

Ica

t (

A)

Nitrite (M)

A

B

Figure 3.20: Variation of Icat with nitrite concentration of ccNiR immobilized on sol-gelfilms A) ccNiR entrapped in a sodium silicate and PEG6000 film B) ccNiR entrapped in asodium silicate and PEG400 film.

The sensitivity of the bioelectrodes, as determined by the slope of the linear part ofthe calibration curve (Figure 3.20) is 197 mA/M.cm2 and 12.7 mA/M.cm2 for the cc-NiR/PEG400 and ccNiR/PEG6000 bioelectrodes, respectively, while the maximum cur-rents at saturating nitrite concentrations (500µM) are -0.109 µA and -2.47 µA, respectively.These results are somewhat contradictory when compared with the non-turnover signalsseen in section 3.2.1)3.2.1.1), considering that ccNiR/PEG6000 gave a poor non- catalyticresponse and in the presence of nitrite, the catalytic response is more than ten-fold thatof ccNiR/PEG400 electrode. Nonetheless, this result was anticipated because, as men-tioned previously, a higher pore size was expected to be more suitable not only for theaccommodation of the large ccNiR aggregates, as well as to guide protein molecules to-wards the electrode surface with PEG6000 based sol- gel., but also for the mass transportof the substrate. Another possibility for the improved behavior of PEG6000 lies on theassumption previously made, that the cathodic peak’s potential is closer to that of thecatalytic site. Within the 3Dmatrix, the sensitivity of the PEG6000 electrode is slightlylower than most of the values reported for the preceding nitrite biosensors constructedwith the same enzyme 39(cf. Table 1.5).

The lower detection limit at a signal to noise ratio of 3 was 2 µM and 0.19 µM forccNiR/PEG400 and ccNiR/PEG6000 bioelectrodes respectively; these values are similarto the previously reported LODs of ccNiR based biosensors and meet the requirementsfor nitrite monitoring in drinking waters according to European Union rules (maximumadmissible level <0.1 ppm, i.e., 2.2 µM). Nevertheless, one should consider further opti-mization studies, using different amounts of enzyme and sol-gel concentrations.

71


3.2.1.3 Stability

The sol-gel/ccNiR modified electrodes were stored at 4Cand tested for stability. Calibra-tion curves were periodically traced and the maximum currents and sensitivity for nitritewere evaluated. Figure 3.21 shows the gradual decrease in the ccNiR’s catalytic responseto nitrite over time (80 days) in a ccNiR/PEG6000 bioelectrode. The ccNiR/PEG400 elec-trodes had a similar profile but maintained their catalytic activity for a shorter period (9days).

-5 0 5 10 15 20 25 30 35 40 45 50 55 60 65 70 75 80 85

-0.010

-0.012

-0.014

-0.016

-0.018

-0.020

Time (days)

Se

nsitiv

ity (A

/M

)

0.0

-0.5

-1.0

-1.5

-2.0

-2.5

-3.0

Ma

x (A

)

0 100 200 300 400 500 600

0.0

-0.2

-0.4

-0.6

-0.8

-1.0

-1.2

Ica

t (

A)

Concentration (M)

A B

Figure 3.21: Variation of Icat with nitrite concentration of ccNiR immobilized on sol-gelfilms A) ccNiR entrapped in a sodium silicate and PEG6000 film B) ccNiR entrapped in asodium silicate and PEG400 film.A) Variation of Icat with nitrite concentration of ccNiRimmobilized on PEG6000 sol-gel films over time. B) Time effects on the biosensor sensi-tivity for nitrite determination. Sensitivity values were given by the slope of calibrationcurves performed periodically throughout 80 days. Catalytic currents wer e measured at- 0.8 V vs Ag/AgCl.

A steep decrease in maximum current and sensitivity occurs in the first day, withboth matrices. Nevertheless, it is clear that the PEG400 leads to a complete loss of ac-tivity faster than the PEG6000 sensor (ccNiR/PEG400 loses activity after 9 days whileccNiR/PEG6000 keeps 50% of its initial activity after 2 weeks). Most likely this is due todenaturation or/and lixiviation of the enzyme in the sol-gel matrices. Though, it is worthmentioning that the long period in which the ccNiR/PEG6000 configuration remains ac-tive is quite promising, and a future optimized formulation possibly comprising enzymestabilizing agents could originate a more reliable and stable biosensor.

72


3.2.2 Macroporous carbon nanotubes

The electrophoretic deposition allows the formation of macroporous CNTs layers fromsuspensions of charged particles on the electrode surface. The carbon nanotubes are neg-atively charged in a broad pH range198 due to the presence of carboxylic groups ontheir surface. They move to the anode during the deposition and form a dense intercon-nected network with a two-dimensional geometry and irregular mesopores caused byvoids between the randomly distributed carbon nanotubes199.The polystyrene beads arealso negatively charged due to the presence of sulfonate groups on their surface (potas-sium persulfate was used as initiator of the polymerization in their synthesis200) and canbe electrophoretically deposited as well. Variation of the deposition time affects thick-ness/amount of CNTs deposited.

The influence of the amount of deposited macroporous SWCNT on the catalytic re-sponse of ccNiR was evaluated by CV on glassy carbon plates. The deposition time wasvaried between 30s and 240s. Figure 3.22 depicts the effect of increasing nitrite concen-trations on the ccNiR/CNT-PS electrode prepared with a 240s deposition time.

-0.9 -0.8 -0.7 -0.6 -0.5 -0.4 -0.3 -0.2 -0.1 0.0

-45

-40

-35

-30

-25

-20

-15

-10

-5

0

5

Cu

rre

nt (

A)


Nitrite

0

3 mM

Figure 3.22: Electrochemical response of PG/CYP1A2–sol-gel film electrode to varyingnitrite concentrations (0-300mM) in 0.1 M KCl and 50 mM tris-HCl buffer pH 7.6 purgedwith argon. Scan rate, 20 mV/s.

Figure 3.23 shows that higher deposition times, and consequent higher nanotubesamounts deposited on the electrode, results in an increased ccNiR response to nitrite.Yet, the sensitivity, as determined by the slope in the linear part of the catalytic currentvs. substrate curve, did not improved with the amount of macroporous material (resultsnot shown) and only a higher maximum current at saturating substrate concentrations

73


was observed.

0 500 1000 1500 2000 2500 3000 3500

0

-10

-20

-30

-40

-50

-60

Ica

t (

A)

[NO2] (M)

Figure 3.23: Catalytic current variation of the ccNir/CNT-PS layer as a function of nitriteconcentration. Macroporous SWCNTs deposition time. (black) 30s; (red) 135s; (green)180s; (blue) 240s.

The deposition times were chosen based on the work developed at LCPM-CNRSNancy201 , which demonstrated that above 240s the film stability starts to decrease. In theconditions tested in this work it was also possible to observe that there is hardly any dif-ference between the 180s and 240s configurations. This behavior is in agreement with theresults presented by Etienne et al.201 very recently, because the film thickness is expectedto start stabilizing after 100s deposition time. Bearing in mind that there is an improvedresponse with the amount of nanotubes, the results presented above are considered pos-itive. Still, these experiments are preliminary and further work must be accomplished tobetter understand the system,for instance, the study of difference between the presenceand absence of the PS beads would be quite interesting.

74


3.2.3 Oxygen scavenger system

The majority of electrochemical nitrite biosensors are used under anaerobic conditions,usually by eliminating the electrolyte’s oxygen content by inert gas bubbling. Herein anenzyme-catalyzed O2 removal system, based on the protocol developed by Plumeré etal.202 will be studied, in order to use a nitrite reductase based biosensor under ambientair electrochemical cells. The glucose-GOx-CAT system (Figure 3.24) was thus tested withrespect to its ability to remove oxygen and maintain anaerobic conditions in unstirredelectrochemical cells open to ambient air.

Figure 3.24: Scheme of the GOx-CAT scavenging system.

The oxygen scavenger system was tested with the sol-gel/ccNiR system describedin section 3.1.1.2)b). In absence of GOx and CAT, the cathodic oxygen reduction signalappears at potentials lower than 600 mV vs Ag/AgCl at the PGE.

Figure 3.25 shows that after addition of glucose, GOx and CAT to the electrochemicalcell oxygen is efficiently removed from the electrolyte. The remaining background cur-rent is comparable or even lower than the one obtained after purging the electrolyte withinert gas for 20 min (usual method for oxygen removal).

75


-0.9 -0.8 -0.7 -0.6 -0.5 -0.4 -0.3 -0.2 -0.1 0.0

-3.50

-3.00

-2.50

-2.00

-1.50

-1.00

-0.50

0.00

0.50

Cu

rre

nt

(A

)


a

b

Figure 3.25: Cyclic voltammograms of PGE/ccNiR/PEG at 20 mV s -1 in 10 mL of 0.1 MKCl and tris-HCl 50 mM pH 7.6; (a) GOx (12.5 µM, 15 UmL -1 ) and CAT (16.6 µM, 2 kUmL -1 ) in solution (b) GOx (12.5 µM, 15 UmL-1), CAT (16.6 µM, 2 kU mL-1) and glucose(50 mM) in solution.

Next, the addition of nitrite to the cell reveals an increase in the catalytic current ofccNiR, indicating that the nitrite biosensor remains functional and is able to detect nitritein the presence of the oxygen scavenging system. (Figure 3.26).

-0.9 -0.8 -0.7 -0.6 -0.5 -0.4 -0.3 -0.2 -0.1 0.0

-0.60

-0.55

-0.50

-0.45

-0.40

-0.35

-0.30

-0.25

-0.20

-0.15

-0.10

-0.05

0.00

0.05

0.10

Cu

rre

nt

(A

)


a

b

Figure 3.26: Cyclic voltammogram with PGE at 20 mV s -1 in 10 mL of 0.1 M KCl and tris-HCl 50 mM pH 7.6; (a) upon addition of GOx (12.5 µM, 15 UmL -1 ) and CAT (16.6 µM, 2kU mL -1 ) and with the addition of glucose (50 mM).(b) addition of 100 mM Nitrite.

76


After confirming the GOx-Catalase-glucose ability to remove oxygen from solution,the efforts were focused on immobilizing the oxygen removal system on the ccNiR bio-electrode in order to constitute a fully integrated biosensor. As a first attempt, catalasewas immobilized on the sol-gel matrix and GOx and glucose were added to the unpurgedelectrolyte. As can be observed in Figure 3.27, this system is not effective for oxygenremoval, since the cathodic oxygen currents are still present. Most likely, catalase is notactive in the sol-gel film or the amount of immobilized enzyme is not enough to completethe cycle of oxygen removal in the vicinity of the electrode surface. Next, the electrolytesolution was thoroughly purged with argon to remove oxygen from solution, howeveras confirmed by the green curve in Figure 3.27A the cathodic currents are not eliminated.At this point it is very likely that the GOx (and glucose in solution) has produced highamounts of hydrogen peroxide which, due to the lack of catalase in solution, is accu-mulating and originating the cathodic currents observed after purging the solution. Inorder to remove the accumulated peroxide, catalase was then added to the solution and,in effect, the cathodic currents were eliminated. Interestingly, upon nitrite addition to thecell, no nitrite reduction currents were observed, which could indicate that the hydrogenperoxide may have inactivated ccNiR.

-0.9 -0.8 -0.7 -0.6 -0.5 -0.4 -0.3 -0.2 -0.1 0.0

-1.0

-0.8

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-0.4

-0.2

0.0

0.2

-0.9 -0.8 -0.7 -0.6 -0.5 -0.4 -0.3 -0.2 -0.1 0.0

-8.0

-6.0

-4.0

-2.0

0.0

Cur

rent

(A

)

E versus Ag/AgCl (V) E versus Ag/AgCl (V)

7

Figure 3.27: Cyclic voltammogram with ccNiR/CAT immobilized in PGE at 20 mV s−1

in 10 mL of 0.1 M KCl and tris-HCl 50 mM pH 7.6; A) ( black ) addition of GOx (12.5 µM,15 UmL−1 ) and of glucose (50 mM).(green) after purging the electrolyte solution withargon for 10 min (magenta) CAT (16.6 µM, 2 kU mL−1 ) (blue) 100 mM Nitrite.B) controlexperiments performed in the absence of ccNiR. (Red line) increasing hydrogen peroxideconcentrations (0-3mM); (black line) non purged electrolyte.

77


The precise chemical mechanisms of the heme destruction are probably very com-plex. The most plausible hypothesis is that hydrogen peroxide produced by GOx is theinitiating agent in the degradative mechanism203,204. Studies on hydrogen peroxide-mediated heme degradation have implicated the hydrogen peroxide complex of ferryl(Fe4+) heme, the ferryl heme itself being produced from a peroxide complex of ferri-and ferrohemes. The peroxide complex of the ferryl heme readily autoxidizes to formthe highly reactive superoxide radical205,206. The proposed mechanism via initially coor-dinated peroxide is consistent with the observations of a lack of reactivity of the hemegroup of ccNiR. Although there is still a lot of work to be accomplished regarding thesystem’s immobilization, the results obtained are very promising from the oxygen scav-enging system point-of-view. It must be taken into account that the phenomenon of hemedegradation by hydrogen peroxide has yet to be reported with ccNiR.

78

4Conclusion

4.1 Cytochrome P450 electrochemistry

The electrochemistry of P450 proteins is rather challenging. Several experimental ap-proaches were tested in order to study this system, ranging from mediated electrochem-istry with solution and electropolymerized electron shuttles, to direct electron transferbased systems using surfactants and/or sol-gel films. However, only a few of thesemethodologies were successful in channeling the electrons from the electrode to the CYP,namely, entrapment with long-chained surfactants, DDM and CTAB, and sodium sili-cate/PEG400 sol-gels.

In this work, P450 DET in a water-based sol-gel thin film is described for the first time.The electrochemistry of the heme center and the electrocatalytic reaction with O2 wasgreatly improved in the presence of PEG400. From the wide range of film compositionstested, it was shown that only the combination of the sodium silicate inorganic matrixand the PEG additive enabled the direct electron transfer reaction and the electrocatalyticactivity towards oxygen.

From this thesis, it becomes apparent that the amount of oxygen in the supportingelectrolyte is critical to the PFeIII/PFeII conversion being monitored on the electrode. Inanaerobic conditions, a small non-reversible cathodic peak is observed, whose intensityincreases with the oxygen concentration. On the other hand, in aerobic conditions, a ca-thodic peak with a high current value is initially observed, which completely disappearsover time. This suggests that the PFeII species is not being regenerated and subsequentlyelectrochemically reduced.

The results obtained with CYP and CYPOR immobilized in sol-gel, in the presenceof varying oxygen and caffeine concentrations raised several questions for which there is

79

4. CONCLUSION 4.2. Cytochrome c nitrite reductase electrochemistry

not a definite answer - why is there no increase in the catalytic current in the presence ofcaffeine? What is the hydrogen’s peroxide relevance for the CYP’s activity? - nonethelessone thing is clear, the CYP’s catalytic cycle is interrupted in the absence of CPR. One of thefirst hypothesis revolved on the prerogative that hydrogen peroxide might have a detri-mental effect on the catalysis of oxygen by CYP, by promoting heme degradation. How-ever, there was no improvement in the system’s response in the presence of a peroxidase(catalase), from which it is possible to infer that if the hydrogen peroxide concentrationwas one of the issues, it certainly was not the only one. The next hypothesis - CYP needsan efficient electron donor, in order to proceed with the substrates conversion, as wellas a specific conformation that may be induced by the interaction with CPR – was sus-tained by the results. In fact, in the presence of CPR a catalytic response to caffeine wasobserved. Most likely, the docking between the two partners (CYP and CPR) is necessaryfor CYP to adopt the right conformation. One the other hand, one should not discardthe hypothesis that the stabilizing matrix is not fully biocompatible, hindering the CYP’scatalytic activity. A decrease in the oxygen catalytic current was observed upon caffeineaddition, indicating that its concentration is being translated into a variation in the oxy-gen response by, somehow, disturbing the CYP cycle. In contrast, Rusling reported anincrease in the catalytic current while using the same P450 isoform and another substrate(styrene)187, regardless in his work it is not mentioned if the substrate stock solution waspurged prior to the addition.

4.2 Cytochrome c nitrite reductase electrochemistry

Aiming at the development/optimization of a fully integrated amperometric nitrite biosen-sor, the hybrid sol-gel used in the studies performed with CYP1A2 was optimized for cc-NiR. The results observed with different sol-gel formulations are in accordance with theones previously obtained with ccNiR, which indicate that large pore matrices are prefer-able for the enzyme’s immobilization/activity124. Indeed, the PEG6000 polymer (ex-pected to produce large pore films) is more favorable than PEG400 to maintain the elec-trocatalysis of nitrite over time. Also, a more favorable orientation might be attained inthis sol-gel, as corroborated by the fact that the cathodic peak’s potential is similar to theone reported for the catalytic site of ccNir as immobilized on a PG electrode207. Macro-porous carbon nanotube assemblies have been successfully prepared by electrophoreticdeposition in the presence of a polystyrene template. The quantity of carbon nanotubesand polystyrene beads in the deposition medium is critical for the proper self-assemblyof the composite film in the 60 V potential field. Cytochrome c nitrite reductase wasdeposited on the surface of these macroporous CNT electrodes and then used for theelectrocatalytic detection of nitrite. A direct correlation was observed between the de-position time and the ccNiR’s electrochemical response to the substrate, as shown byan increase in the catalytic current with nitrite concentration. Most likely, the greater

80


nanotube surface area generated with longer deposition times allows the electrical con-nection of a higher amount of electroactive enzyme. There are still other venues thathave to be followed in order to ascertain the real weight of this approach, namely theinterference with the size of the PS beads in the ccNiR’s response to substrate. An oxy-gen removal system based in an oxidase, in the presence of its aldohexose substrate, andcatalase was investigated for field biosensor applications. Electrochemical measurementsunder ambient air were free of oxygen interferences, even for electrolyte volumes of 10mL. Quantification of nitrite, in an open cell with an electrochemical biosensor based onccNiR as the biorecognition element and GOx as O2 reduction catalyst was demonstrated.Interestingly, the results obtained during the study of this oxygen scavenger system - thebioelectrode lost its activity after being exposed to a high concentration of peroxide -uphold previous theories postulating that hydrogen peroxide might contribute to hemegroup degradation208. Although, thus far it was not possible to immobilize the oxygenscavenger system, this approach was successful in maintaining anaerobic conditions inthe model nitrite biosensor described, proving itself a promising, simple and inexpensiveoxygen removal methodology.

81


82

5Future work

5.1 Cytochrome P450 electrochemistry

Several authors have proposed that any electrochemical experiments of immobilizedP450 should be accompanied by control experiments with P420 (the inactive form), aswell as an iron-protoprofirin group or other heme proteins such as myoglobin. Thereforeas future work we propose to repeat this experiments using two working electrodes, onewith immobilized P450 and the other with immobilized P420. Voltammograms shouldthen be measured under identical conditions by switching between the two working elec-trodes. This would allow us to assure that the P450 has not been inactivated and detectother possible electrochemical artefacts. Another interesting experiment would be to co-immobilized catalase with P450 and P420 with the aim of reducing or eliminating hydro-gen peroxide.

Additionally, other immobilization matrices should be tested with the CYP/CYPORsystem in order to improve the electrochemical response and catalysis. Furthermore,other P450 substrates should be tested, for example propanolol and paracetamol.

Seeing that, the electron tunneling with DET is not mimicking the physiological elec-tron pathway, future parallel studies could be performed with different redox mediatorsand electrochemical potential windows in order to access CYP’s activity.

5.2 Cytochrome c nitrite reductase electrochemistry

The hybrid sol-gel developed in this thesis is a very promising biocompatible matrix,mainly due to the fact that is a water based formulation. Therefore, future work shouldstrive to improve this matrix, for instance for application in nitrite biosensors. These

83

5. FUTURE WORK 5.2. Cytochrome c nitrite reductase electrochemistry

attempts should comprise further optimization of the sodium silicate, PEG and ccNiRratios in the sol-gel film, the use of different molecular weight PEGs or the incorporationof other sol-gel stabilizing agents, such as PEI. It would also be quite interesting to per-form surface characterization, of these bioelectrodes, in order to confirm effective enzymeimmobilization, by scanning electron microscopy or atomic force microscopy.

Regarding the macroporous nanotubes electrode modification system, future endeav-ors should cover a more detailed study of the importance of the CNT’s film porosity. Thatbeing said, the GC-plates should be functionalized with varying deposition times withand without PS-beads.

In the future, whichever sensor platforms used, it will be beneficial to promote theenzyme molecules structural orientation on the bioelectrodes, which would guarantee agreater reproducibility in the amount of electrically-wired enzyme and, consequently, thebiosensor’s response. This could be achieved by the use of functionalized materials, suchas self-assembled monolayers or gold nanoparticles.

In order to implement the enzymatic oxygen scavenger system in biosensors, its im-mobilization must be accomplished; the sol-gel matrix here described should be opti-mized for this purpose or a different encapsulation material could be tested.

84

6References

1. Borgmann, S., Schulte, A., Neugebauer, S. & Schuhmann, W. Amperometric Biosen-sors. Advances in Electrochemical Science and Engineering: Bioelectrochemistry(2011).doi:10.1002/9783527644117.ch1

2. Urban, G. Reinhard Renneberg and Fred Lisdat (eds.): Biosensing for the 21st cen-tury. Analytical and Bioanalytical Chemistry 393, 777-778 (2008).

3. Zweier, J. L., Li, H., Samouilov, A. & Liu, X. Mechanisms of nitrite reduction tonitric oxide in the heart and vessel wall. Nitric oxide: biology and chemistry /official journal of the Nitric Oxide Society 22, 83-90 (2010).

4. Severinghaus, J. W. The invention and development of blood gas analysis appara-tus. Anesthesiology 97, 253–6 (2002).

5. Clark, L. C. & Lyons, C. Electrode systems for continuous monitoring in cardiovas-cular surgery. Annals of the New York Academy of Sciences 102, 29–45 (2006).

6. Wang, J. Electrochemical glucose biosensors. Chemical reviews 108, 814–25 (2008).

7. Almeida, M. G. Electrochemical biosensors revisited: old solutions to new prob-lems. Submited in Bioelectrochemistry (2013).

8. Guilbault, G. G. & Lubrano, G. J. An enzyme electrode for the amperometric deter-mination of glucose. Analytica chimica acta 64, 439–55 (1973).

9. Cammann, K. Bio-sensors based on ion-selective electrodes. Fresenius’ Zeitschriftfor Analytische Chemie 287, 1–9 (1977).

85

6. REFERENCES

10. Eddowes, M. J. & Hill, H. A. O. Novel method for the investigation of the elec-trochemistry of metalloproteins: cytochrome c. Journal of the Chemical Society,Chemical Communications 771b (1977).doi:10.1039/c3977000771b

11. Eddowes, M. J., Hill, H. A. O. & Uosaki, K. The electrochemistry of cytochromec. Investigation of the mechanism of the 4,4’-bipyridyl surface modified gold elec-trode. Journal of Electroanalytical Chemistry and Interfacial Electrochemistry 116,527–537 (1980).

12. Huck, H. & Schmidt, H.-L. Chloranil as a Catalyst for the Electrochemical Oxidationof NADH to NAD+. Angewandte Chemie International Edition in English 20, 402–403 (1981).

13. Jaegfeldt, H., Torstensson, A. B. C., Gorton, L. G. O. & Johansson, G. Catalytic oxida-tion of reduced nicotinamide adenine dinucleotide by graphite electrodes modifiedwith adsorbed aromatics containing catechol functionalities. Analytical Chemistry53, 1979–1982 (1981).

14. Shichiri, M., Yamasaki, Y., Kawamori, R., Hakui, N. & Abe, H. Wearable artificialendocrine pancreas with needle-type glucose sensor. The Lancet 320, 1129–1131(1982).

15. Cass, A. E. G. et al. Ferrocene-mediated enzyme electrode for amperometric deter-mination of glucose. Analytical Chemistry 56, 667–671 (1984).

16. Degani, Y. & Heller, A. Direct electrical communication between chemically mod-ified enzymes and metal electrodes. I. Electron transfer from glucose oxidase tometal electrodes via electron relays, bound covalently to the enzyme. The Journalof Physical Chemistry 91, 1285–1289 (1987).

17. Degani, Y. & Heller, A. Direct electrical communication between chemically modi-fied enzymes and metal electrodes. 2. Methods for bonding electron-transfer relaysto glucose oxidase and D-amino-acid oxidase. Journal of the American ChemicalSociety 110, 2615–2620 (1988).

18. Forster, R. J. & Vos, J. G. Synthesis, characterization, and properties of a series ofosmium- and ruthenium-containing metallopolymers. Macromolecules 23, 4372–4377 (1990).

19. Csöregi, E., Schmidtke, D. W. & Heller, A. Design and optimization of a selectivesubcutaneously implantable glucose electrode based on “wired” glucose oxidase.Analytical chemistry 67, 1240–4 (1995).

20. Schmidtke, D. W. Measurement and modeling of the transient difference betweenblood and subcutaneous glucose concentrations in the rat after injection of insulin.Proceedings of the National Academy of Sciences 95, 294–299 (1998).

86

6. REFERENCES

21. Wagner, J. G. et al. Continuous amperometric monitoring of glucose in a brittlediabetic chimpanzee with a miniature subcutaneous electrode. Proceedings of theNational Academy of Sciences 95, 6379–6382 (1998).

22. Frew, J. E. & Hill, H. a Direct and indirect electron transfer between electrodes andredox proteins. European journal of biochemistry / FEBS 172, 261–9 (1988).

23. Tarasevich, M. R., Yaropolov, A. I., Bogdanovskaya, V. A. & Varfolomeev, S. D. 293 -Electrocatalysis of a cathodic oxygen reduction by laccase. Bioelectrochemistry andBioenergetics 6, 393–403 (1979).

24. IKEDA, T., FUSHIMI, F., MIKI, K. & SENDA, M. Direct bioelectrocatalysis at elec-trodes modified with D-gluconate dehydrogenase. Agricultural and biological chem-istry 52, 2655–2658.

25. Ikeda, T., Miyaoka, S. & Miki, K. Enzyme-catalysed electrochemical oxidation ofd-gluconate at electrodes coated with d-gluconate dehydrogenase, a membrane-bound flavohemoprotein. Journal of Electroanalytical Chemistry 352, 267–278 (1993).

26. Armstrong, F. A., George, S. J., Thomson, A. J. & Yates, M. G. Direct electrochem-istry in the characterisation of redox proteins: Novel properties of Azotobacter 7Feferredoxin. FEBS Letters 234, 107–110 (1988).

27. Armstrong, F. A. Probing Metalloproteins by Voltammetry. Bioinorganic Chemistry72, (1990).

28. Armstrong, F. A., Heering, H. A. & Hirst, J. Reaction of complex metalloproteinsstudied by protein-film voltammetry. Chemical Society Reviews 26, 169 (1997).

29. Zayats, M., Kharitonov, A. B., Katz, E., Bückmann, A. F. & Willner, I. An integratedNAD+-dependent enzyme-functionalized field-effect transistor (ENFET) system:development of a lactate biosensor. Biosensors & bioelectronics 15, 671–80 (2000).

30. Kurzawa, C., Hengstenberg, A. & Schuhmann, W. Immobilization method for thepreparation of biosensors based on pH shift-induced deposition of biomolecule-containing polymer films. Analytical chemistry 74, 355–61 (2002).

31. Guschin, D. a, Shkil, H. & Schuhmann, W. Electrodeposition polymers as immo-bilization matrices in amperometric biosensors: improved polymer synthesis andbiosensor fabrication. Analytical and bioanalytical chemistry 395, 1693–706 (2009).

32. Guschin, D. a, Castillo, J., Dimcheva, N. & Schuhmann, W. Redox electrodeposi-tion polymers: adaptation of the redox potential of polymer-bound Os complexesfor bioanalytical applications. Analytical and bioanalytical chemistry 398, 1661–73(2010).

87

6. REFERENCES

33. Weinstein, R. L. et al. Accuracy of the 5-day FreeStyle Navigator Continuous Glu-cose Monitoring System: comparison with frequent laboratory reference measure-ments. Diabetes care 30, 1125–30 (2007).

34. Scheller, Frieder W.; Schubert, F. et al. Biosensors. Journal of Chromatography A510, 347–354 (Elsevier: 1992).

35. Thévenot, D. R., Toth, K., Durst, R. A., Wilson, G. S. & The, D. R. Electrochemicalbiosensors: recommended definitions and classification. Biosensors and Bioelec-tronics 16, 121–131 (2001).

36. Cracknell, J. a, Vincent, K. a & Armstrong, F. a Enzymes as working or inspirationalelectrocatalysts for fuel cells and electrolysis. Chemical reviews 108, 2439–61 (2008).

37. Guilbault, G. G. Use of immobilized enzymes in chemical analysis. Journal of Solid-Phase Biochemistry 2, 329–342 (1978).

38. Phillips, P. E. M. & Wightman, R. M. Critical guidelines for validation of the selec-tivity of in-vivo chemical microsensors. TrAC Trends in Analytical Chemistry 22,509–514 (2003).

39. Silveira, C. M. C. F. Development of electrochemical nitrite biosensors using cy-tochrome c nitrite reductase from Desulfovibrio desulfuricans ATCC 27774. (2011).at<http://hdl.handle.net/10362/6811>

40. Wang, Z., Etienne, M., Quilès, F., Kohring, G.-W. & Walcarius, A. Durable cofactorimmobilization in sol-gel bio-composite thin films for reagentless biosensors andbioreactors using dehydrogenases. Biosensors & bioelectronics 32, 111–7 (2012).

41. Fernandez-lafuente, R. et al. Stabilization of multimeric enzymes via immobiliza-tion and post-immobilization techniques. Journal of Molecular Catalysis B: Enzy-matic 7, 181–189 (1999).

42. Torchilin, V. P., Trubetskoy, V. S., Omel’Yanenko, V. G. & Mratinek, K. Stabilizationof subunit enzyme by intersubunit crosslinking with bifunctional reagents: studieswith glyceraldehyde-3-phosphate dehydrogenase. Journal of Molecular Catalysis19, 291–301 (1983).

43. Scouten, W., Luong, J. & Stephenbrown, R. Enzyme or protein immobilization tech-niques for applications in biosensor design. Trends in Biotechnology 13, 178–185(1995).

44. Vandenberg, E., Brown, R. & Krull, U. Immobilized biosystems in theory and prac-tical applications. Food / Nahrung 39, 129 (Elsevier, Holland: 1983).

88

6. REFERENCES

45. Weetall, H. H. Preparation of immobilized proteins covalently coupled throughsilane coupling agents to inorganic supports. Applied biochemistry and biotech-nology 41, 157–88 (1993).

46. Doretti, L., Ferrara, D. & Lora, S. Enzyme-entrapping membranes for biosensorsobtained by radiation-induced polymerization. Biosensors and Bioelectronics 8,443–450 (1993).

47. O’Driscoll, K. F. Techniques of enzyme entrapment in gels. Methods in enzymology44, 169–83 (1976).

48. Scouten, W. H. A survey of enzyme coupling techniques. Immobilized Enzymesand Cells Part B 135, 30–60 (1987).

49. Adeyoju, O., Iwuoha, E. I., Smyth, M. R. & Leech, D. High-performance liquidchromatographic determination of phenols using a tyrosinase-based amperometricbiosensor detection system. The Analyst 121, 1885 (1996).

50. Scouten, W. H., Luong, J. H. T., Stephenbrown, R. & Brown, R. S. Enzyme or proteinimmobilization techniques for applications in biosensor design. Trends in Biotech-nology 13, 178–185 (1995).

51. Chibata, I. Immobilized enzymes — research and development. Biochemical Edu-cation 7, 284 (Wiley & Sons, Incorporated, John: 1978).

52. Avnir, D., Coradin, T., Lev, O. & Livage, J. Recent bio-applications of sol–gel mate-rials. Journal of Materials Chemistry 16, 1013 (2006).

53. Wang, J. Sol–gel materials for electrochemical biosensors. Analytica Chimica Acta399, 21–27 (1999).

54. Walcarius, A. & Collinson, M. M. Analytical chemistry with silica sol-gels: tradi-tional routes to new materials for chemical analysis. Annual review of analyticalchemistry (Palo Alto, Calif.) 2, 121–43 (2009).

55. Ellerby, L. et al. Encapsulation of proteins in transparent porous silicate glassesprepared by the sol-gel method. Science 255, 1113–1115 (1992).

56. Gill, I. & Ballesteros, A. Bioencapsulation within synthetic polymers (Part 2): non-sol-gel protein-polymer biocomposites. Trends in biotechnology 18, 469–79 (2000).

57. Pastor, I., Prieto, M. & Mateo, C. R. Effect of sol-gel confinement on the structuraldynamics of the enzyme bovine Cu,Zn superoxide dismutase. The journal of phys-ical chemistry. B 112, 15021–8 (2008).

58. Jin, W. & Brennan, J. D. Properties and applications of proteins encapsulated withinsol–gel derived materials. Analytica Chimica Acta 461, 1–36 (2002).

89

6. REFERENCES

59. Seddon, A. M., Curnow, P. & Booth, P. J. Membrane proteins, lipids and detergents:not just a soap opera. Biochimica et biophysica acta 1666, 105–17 (2004).

60. Le Maire, M., Champeil, P. & Moller, J. V Interaction of membrane proteins andlipids with solubilizing detergents. Biochimica et biophysica acta 1508, 86–111(2000).

61. Vittal, R., Gomathi, H. & Kim, K.-J. Beneficial role of surfactants in electrochemistryand in the modification of electrodes. Advances in colloid and interface science 119,55–68 (2006).

62. Rusling, J. J. F. Enzyme bioelectrochemistry in cast biomembrane-like films. Ac-counts of chemical research 31, 363–369 (1998).

63. Nassar, A. E., Willis, W. S. & Rusling, J. F. Electron transfer from electrodes to myo-globin: facilitated in surfactant films and blocked by adsorbed biomacromolecules.Analytical chemistry 67, 2386–92 (1995).

64. Nassar, A. E., Rusling, J. F. & Kumosinski, T. F. Salt and pH effects on electrochem-istry of myoglobin in thick films of a bilayer-forming surfactant. Biophysical chem-istry 67, 107–16 (1997).

65. Lu, Z., Huang, Q. & Rusling, J. F. Films of hemoglobin and didodecyldimethylam-monium bromide with enhanced electron transfer rates. Journal of ElectroanalyticalChemistry 423, 59–66 (1997).

66. Mimica, D., Zagal, J. H. & Bedioui, F. Electroreduction of nitrite by hemin, myo-globin and hemoglobin in surfactant films. Journal of Electroanalytical Chemistry497, 106–113 (2001).

67. Guto, P. M. & Rusling, J. F. Myoglobin retains iron heme and near-native conforma-tion in DDAB films prepared from pH 5 to 7 dispersions. Electrochemistry Com-munications 8, 455–459 (2006).

68. Rusling, J. F. & Nassar, A. E. F. Enhanced electron transfer for myoglobin in surfac-tant films on electrodes. Journal of the American Chemical Society 115, 11891–11897(1993).

69. Zhang, Z., Nassar, A. F., Lu, Z., Schenkman, J. B. & Rusling, J. F. Direct electron in-jection from electrodes to cytochrome P450cam in biomembrane-like films. Journalof the Chemical Society, Faraday Transactions 93, 1769–1774 (1997).

70. Nassar, A.-E. F., Zhang, Z., Hu, N., Rusling, J. F. & Kumosinski, T. F. Proton-CoupledElectron Transfer from Electrodes to Myoglobin in Ordered Biomembrane-like Films.The Journal of Physical Chemistry B 101, 2224–2231 (1997).

90

6. REFERENCES

71. Zu, X., Lu, Z., Zhang, Z., Schenkman, J. B. & Rusling, J. F. Electroenzyme-CatalyzedOxidation of Styrene and cis- -Methylstyrene Using Thin Films of Cytochrome P450camand Myoglobin. 7372–7377 (1999).

72. Johnson, D. L., Lewis, B. C., Elliot, D. J., Miners, J. O. & Martin, L. L. Electrochemicalcharacterisation of the human cytochrome P450 CYP2C9. Biochemical pharmacol-ogy 69, 1533–41 (2005).

73. Udit, A. K., Hindoyan, N., Hill, M. G., Arnold, F. H. & Gray, H. B. Protein-surfactantfilm voltammetry of wild-type and mutant cytochrome P450 BM3. Inorganic chem-istry 44, 4109–11 (2005).

74. Aguey-Zinsou, K.-F., Bernhardt, P. V, De Voss, J. J. & Slessor, K. E. Electrochemistryof P450cin: new insights into P450 electron transfer. Chemical communications(Cambridge, England) 418–9 (2003).at <http://www.ncbi.nlm.nih.gov/pubmed/12613641>

75. Shukla, A., Gillam, E. M., Mitchell, D. J. & Bernhardt, P. V Direct electrochemistry ofenzymes from the cytochrome P450 2C family. Electrochemistry Communications7, 437–442 (2005).

76. Marcus, R. A. & Sutin, N. Electron transfers in chemistry and biology. Biochimicaet Biophysica Acta (BBA) - Reviews on Bioenergetics 811, 265–322 (1985).

77. Habermüller, K., Mosbach, M. & Schuhmann, W. Electron-transfer mechanismsin amperometric biosensors. Fresenius’ journal of analytical chemistry 366, 560–8 (2000).

78. Guo, L.-H., Allen, H. & Hill, O. Direct Electrochemistry of Proteins and Enzymes.Advances in Inorganic Chemistry 341–375 (1991).

79. Fridman, V. et al. Electrochemical investigation of cellobiose oxidation by cellobiosedehydrogenase in the presence of cytochrome c as mediator. Biochemical Societytransactions 28, 63–70 (2000).

80. Scheller, F. W. et al. Bioelectrocatalysis by redox enzymes at modified electrodes.Reviews in Molecular Biotechnology 82, 411–424 (2002).

81. Léger, C., Bertrand, P. & Le, C. Direct electrochemistry of redox enzymes as a toolfor mechanistic studies. Chemical reviews 108, 2379–438 (2008).

82. Kinnear, K. T. & Monbouquette, H. G. Direct electron transfer to Escherichia colifumarate reductase in self-assembled alkanethiol monolayers on gold electrodes.Langmuir 9, 2255–2257 (1993).

83. Sucheta, A., Cammack, R., Weiner, J. & Armstrong, F. A. Reversible electrochem-istry of fumarate reductase immobilized on an electrode surface. Direct voltammet-ric observations of redox centers and their participation in rapid catalytic electrontransport. Biochemistry 32, 5455–5465 (1993).

91

6. REFERENCES

84. Sucheta, A., Ackrell, B. A., Cochran, B. & Armstrong, F. A. Diode-like behaviour ofa mitochondrial electron-transport enzyme. Nature 356, 361–2 (1992).

85. Bianco, P. Electrocatalytic Hydrogen-Evolution at the Pyrolytic Graphite Electrodein the Presence of Hydrogenase. Journal of The Electrochemical Society 139, 2428(1992).

86. Khan, G. F., Shinohara, H., Ikariyama, Y. & Aizawa, M. Electrochemical behaviourof monolayer quinoprotein adsorbed on the electrode surface. Journal of Electroan-alytical Chemistry and Interfacial Electrochemistry 315, 263–273 (1991).

87. Ikeda, T., Matsushita, F. & Senda, M. Amperometric fructose sensor based on directbioelectrocatalysis. Biosensors and Bioelectronics 6, 299–304 (1991).

88. Burrows, A. L. et al. Direct electrochemistry of the enzyme, methylamine dehydro-genase, from bacterium W3A1. European journal of biochemistry / FEBS 199, 73–8(1991).

89. Guo, L. H., Hill, H. A., Hopper, D. J., Lawrance, G. A. & Sanghera, G. S. Directvoltammetry of the Chromatium vinosum enzyme, sulfide:cytochrome c oxidore-ductase (flavocytochrome c552). The Journal of biological chemistry 265, 1958–63(1990).

90. Paddock, R. M. & Bowden, E. F. Electrocatalytic reduction of hydrogen peroxide viadirect electron transfer from pyrolytic graphite electrodes to irreversibly adsorbedcytochrome c peroxidase. Journal of Electroanalytical Chemistry and InterfacialElectrochemistry 260, 487–494 (1989).

91. Armstrong, F. A. & Lannon, A. M. Fast interfacial electron transfer between cy-tochrome c peroxidase and graphite electrodes promoted by aminoglycosides: novelelectroenzymic catalysis of hydrogen peroxide reduction. Journal of the AmericanChemical Society 109, 7211–7212 (1987).

92. Guo, L. H., Hill, H. A. O., Lawrance, G. A., Sanghera, G. S. & Hopper, D. J. Directun-mediated electrochemistry of the enzyme P-cresolmethylhydroxylase. Journalof Electroanalytical Chemistry and Interfacial Electrochemistry 266, 379–396 (1989).

93. Yaropolov, A. I., Karyakin, A. A., Varfolomeev, S. D. & Berezin, I. V. Mechanismof H2-electrooxidation with immobilized hydrogenase. Bioelectrochemistry andBioenergetics 12, 267–277 (1984).

94. E Ferapontova, E. & Gorton, L. Direct electrochemistry of heme multicofactor-containingenzymes on alkanethiol-modified gold electrodes. Bioelectrochemistry (Amster-dam, Netherlands) 66, 55–63 (2005).

95. Barton, S. C., Gallaway, J. & Atanassov, P. Enzymatic biofuel cells for implantableand microscale devices. Chemical reviews 104, 4867–86 (2004).

92

6. REFERENCES

96. Silveira, C. M. & Almeida, M. G. Small electron-transfer proteins as mediatorsin enzymatic electrochemical biosensors. Analytical and bioanalytical chemistry(2013).doi:10.1007/s00216-013-6786-4

97. Ronkainen, N. J., Halsall, H. B. & Heineman, W. R. Electrochemical biosensors.Chemical Society reviews 39, 1747–63 (2010).

98. Yamamoto, K., Takagi, K., Kano, K. & Ikeda, T. Bioelectrocatalytic Detection of His-tamine Using Quinohemoprotein Amine Dehydrogenase and the Native ElectronAcceptor Cytochrome c-550. Electroanalysis 13, 375–379 (2001).

99. Townshend, A. Frontiers in biosensorics, I. fundamental aspects, II practical appli-cations. Analytica Chimica Acta 353, 398 (1997).

100. Ghindilis, A. L., Atanasov, P. & Wilkins, E. Enzyme-catalyzed direct electron trans-fer: Fundamentals and analytical applications. Electroanalysis 9, 661–674 (1997).

101. Gorton, L. et al. Direct electron transfer between heme-containing enzymes andelectrodes as basis for third generation biosensors. Analytica Chimica Acta 400,91–108 (1999).

102. Updike, S. J. & Hicks, G. P. The Enzyme Electrode. Nature 214, 986–988 (1967).

103. Wollenberger, U., Schubert, F., Pfeiffer, D. & Scheller, F. W. Enhancing biosensor per-formance using multienzyme systems. Trends in biotechnology 11, 255–62 (1993).

104. Spalding, R. F. & Exner, M. E. Occurrence of Nitrate in Groundwater—A Review.Journal of Environment Quality 22, 392 (1993).

105. Manassaram, D. M., Backer, L. C. & Moll, D. M. A Review of Nitrates in Drink-ing Water: Maternal Exposure and Adverse Reproductive and Developmental Out-comes. Environmental Health Perspectives 114, 320–327 (2005).

106. Carpenter, S. R. et al. Nonpoint pollution of surface waters with phosphorous andnitrogen. Ecological Applications 8, 559–568 (1998).

107. Vitousek, P. M. et al. Human alteration of the global nirtrogen cycle: sources andconsequences. Ecological Applications 7, 737–750 (1997).

108. COMLY, H. H. Cyanosis in infants caused by nitrates in well water. JAMA: TheJournal of the American Medical Association 129, 112 (1945).

109. Almeida, G., Tavares, P., Lampreia, J., Moura, J. & Moura, I. Abstracts Poster Pre-sentations (A). Journal of Inorganic Biochemistry 86, 119–130 (2001).

110. Lundberg, J. & Gladwin, M. Nitrate and nitrite in biology, nutrition and therapeu-tics. Nature chemical biology 5, 865–869 (2009).

93

6. REFERENCES

111. Shiva, S., Gladwin, M. T. & Gladwin, Æ. M. T. Nitrite mediates cytoprotection af-ter ischemia/reperfusion by modulating mitochondrial function. Basic research incardiology 104, 113–9 (2009).

112. Hord, N. G., Tang, Y. & Bryan, N. S. Food sources of nitrates and nitrites: the physio-logic context for potential health benefits. The American journal of clinical nutrition90, 1–10 (2009).

113. Van Faassen, E. E. et al. Nitrite as regulator of hypoxic signaling in mammalianphysiology. Medicinal research reviews 29, 683–741 (2009).

114. Bryan, N. S. Nitrite in nitric oxide biology: cause or consequence? A systems-basedreview. Free radical biology & medicine 41, 691–701 (2006).

115. Reynolds, J. D. et al. Nitrate and nitrite anion concentration in the intact cerebralcortex of preterm and nearterm fetal sheep: indirect index of in vivo nitric oxideformation. Journal of pharmacological and toxicological methods 39, 125–8 (1998).

116. Raat, N. J. H. et al. Dietary nitrate and nitrite modulate blood and organ nitriteand the cellular ischemic stress response. Free radical biology & medicine 47, 510–7(2009).

117. Cosby, K. et al. Nitrite reduction to nitric oxide by deoxyhemoglobin vasodilatesthe human circulation. Nature medicine 9, 1498–505 (2003).

118. Sinha, S. S., Shiva, S. & Gladwin, M. T. Myocardial protection by nitrite: evidencethat this reperfusion therapeutic will not be lost in translation. Trends in cardiovas-cular medicine 18, 163–72 (2008).

119. Jensen, F. B. The role of nitrite in nitric oxide homeostasis: a comparative perspec-tive. Biochimica et biophysica acta 1787, 841–8 (2009).

120. Perlman, D. H. et al. Mechanistic insights into nitrite-induced cardioprotection us-ing an integrated metabolomic/proteomic approach. Circulation research 104, 796–804 (2009).

121. Bryan, N. S. et al. Dietary nitrite supplementation protects against myocardialischemia-reperfusion injury. Proceedings of the National Academy of Sciences ofthe United States of America 104, 19144–9 (2007).

122. Webb, A. J. et al. Acute blood pressure lowering, vasoprotective, and antiplateletproperties of dietary nitrate via bioconversion to nitrite. Hypertension 51, 784–90(2008).

123. Ellis, G., Adatia, I., Yazdanpanah, M. & Makela, S. K. Nitrite and nitrate analyses: aclinical biochemistry perspective. Clinical biochemistry 31, 195–220 (1998).

94

6. REFERENCES

124. Silveira, C. M. et al. An efficient non-mediated amperometric biosensor for nitritedetermination. Biosensors & bioelectronics 25, 2026–32 (2010).

125. Almeida, M. G., Serra, A., Silveira, C. M. & Moura, J. J. G. Nitrite biosensing viaselective enzymes–a long but promising route. Sensors (Basel, Switzerland) 10,11530–55 (2010).

126. Bianco, P. The Origin of Bioelectrochemistry: An Overview. Encyclopedia of Elec-trochemistry, Bioelectrochemistry (Wiley-VCH: 2002).

127. Silveira, C. M., Besson, S., Moura, I., Moura, J. J. G. & Almeida, M. G. Measuringthe cytochrome C nitrite reductase activity-practical considerations on the enzymeassays. Bioinorganic chemistry and applications 2010, (2010).

128. Da Silva, S., Cosnier, S., Gabriela Almeida, M. & Moura, J. J. G. An efficient poly(pyrrole–viologen)-nitrite reductase biosensor for the mediated detection of nitrite. Electro-chemistry Communications 6, 404–408 (2004).

129. Serra, a S. et al. Cooperative use of cytochrome cd1 nitrite reductase and its redoxpartner cytochrome c552 to improve the selectivity of nitrite biosensing. Analyticachimica acta 693, 41–6 (2011).

130. Almeida, M. G., Silveira, C. M. & Moura, J. J. G. Biosensing nitrite using the sys-tem nitrite redutase/Nafion/methyl viologen–a voltammetric study. Biosensors &bioelectronics 22, 2485–92 (2007).

131. Kissinger, P. T. Biosensors-a perspective. Biosensors & bioelectronics 20, 2512–6(2005).

132. Chen, H. et al. Highly sensitive nitrite biosensor based on the electrical wiring ofnitrite reductase by [ZnCr-AQS] LDH. Electrochemistry Communications 9, 2240–2245 (2007).

133. Richardson, D. J. & Watmough, N. J. Inorganic nitrogen metabolism in bacteria.Current opinion in chemical biology 3, 207–19 (1999).

134. Moura, I. & Moura, J. J. Structural aspects of denitrifying enzymes. Current opinionin chemical biology 5, 168–75 (2001).

135. Kiang, C. H., Kuan, S. S. & Guilbault, G. G. A novel enzyme electrode method forthe determination of nitrite based on nitrite reductase. Analytica chimica acta 80,209–14 (1975).

136. Scharf, M. et al. Electrochemical Studies on Nitrite Reductase toward a Biosensor.Biochemical and Biophysical Research Communications 209, 1018–1025 (1995).

95

6. REFERENCES

137. Zhang, Z. et al. A novel nitrite biosensor based on conductometric electrode mod-ified with cytochrome c nitrite reductase composite membrane. Biosensors & bio-electronics 24, 1574–9 (2009).

138. Silveira, C. M. et al. Enhanced Direct Electron Transfer of a Multihemic Nitrite Re-ductase on Single-walled Carbon Nanotube Modified Electrodes. Electroanalysis22, 2973–2978 (2010).

139. Strehlitz, B. et al. A nitrite sensor based on a highly sensitive nitrite reductasemediator-coupled amperometric detection. Analytical chemistry 68, 807–16 (1996).

140. Strehlitz, B. et al. Artificial electron donors for nitrate and nitrite reductases usableas mediators in amperometric biosensors. Fresenius’ Journal of Analytical Chem-istry 349, 676–678 (1994).

141. Ferretti, S. et al. Optical biosensing of nitrite ions using cytochrome cd1 nitrite re-ductase encapsulated in a sol-gel matrix. The Analyst 125, 1993–9 (2000).

142. Rosa, C. C., Cruz, H. J., Vidal, M. & Oliva, A. G. Optical biosensor based on nitritereductase immobilised in controlled pore glass. Biosensors & bioelectronics 17, 45–52 (2002).

143. Wu, Q. et al. A nitrite biosensor based on a maltose binding protein nitrite reduc-tase fusion immobilized on an electropolymerized film of a pyrrole-derived bipyri-dinium. Analytical chemistry 69, 4856–63 (1997).

144. Quan, D., Min, D. G., Cha, G. S. & Nam, H. Electrochemical characterization ofbiosensor based on nitrite reductase and methyl viologen co-immobilized glassycarbon electrode. Bioelectrochemistry (Amsterdam, Netherlands) 69, 267–75 (2006).

145. Sasaki, S. et al. Application of nitrite reductase from Alcaligenes faecalis S-6 fornitrite measurement. Biosensors & bioelectronics 13, 1–5 (1998).

146. Astier, Y. et al. Sensing nitrite through a pseudoazurin-nitrite reductase electrontransfer relay. Chemphyschem: a European journal of chemical physics and physi-cal chemistry 6, 1114–20 (2005).

147. Einsle, O. Cytochrome c nitrite reductase. Handbook of metalloproteins 1–14 (2006).doi:10.1002/0470028637.met131

148. Rodrigues, M. L., Oliveira, T. F., Pereira, I. a C. & Archer, M. X-ray structure of themembrane-bound cytochrome c quinol dehydrogenase NrfH reveals novel haemcoordination. The EMBO journal 25, 5951–60 (2006).

149. Almeida, M. G. et al. The isolation and characterization of cytochrome c nitrite re-ductase subunits (NrfA and NrfH) from Desulfovibrio desulfuricans ATCC 27774.European Journal of Biochemistry 270, 3904–3915 (2003).

96

6. REFERENCES

150. Einsle, O. et al. Structure of cytochrome c nitrite reductase. Nature 400, 476–80(1999).

151. Cunha, C. a et al. Cytochrome c nitrite reductase from Desulfovibrio desulfuricansATCC 27774. The relevance of the two calcium sites in the structure of the catalyticsubunit (NrfA). The Journal of biological chemistry 278, 17455–65 (2003).

152. Bamford, V. a et al. Structure and Spectroscopy of the Periplasmic Cytochrome cNitrite Reductase from Escherichia coli. Biochemistry 41, 2921–2931 (2002).

153. Einsle, O. et al. Cytochrome c nitrite reductase from Wolinella succinogenes. Struc-ture at 1.6 A resolution, inhibitor binding, and heme-packing motifs. The Journalof biological chemistry 275, 39608–16 (2000).

154. Bellamine, A., Mangla, A. T., Nes, W. D. & Waterman, M. R. Characterization andcatalytic properties of the sterol 14alpha-demethylase from Mycobacterium tuber-culosis. Proceedings of the National Academy of Sciences of the United States ofAmerica 96, 8937–42 (1999).

155. Shumyantseva, V. V. et al. Electrochemical investigations of cytochrome P450. Biochim-ica et Biophysica Acta (BBA) - Proteins & Proteomics 1814, 94–101 (2011).

156. Montellano, P. R. O. De cytochrome p450: structure mechanism and biochemistry.(Springer-Verlag New York: 1995).

157. Sadeghi, S. J., Fantuzzi, A. & Gilardi, G. Breakthrough in P450 bioelectrochemistryand future perspectives. Biochimica et biophysica acta 1814, 237–248 (2011).

158. Hlavica, P. Assembly of non-natural electron transfer conduits in the cytochromeP450 system: a critical assessment and update of artificial redox constructs amenableto exploitation in biotechnological areas. Biotechnology advances 27, 103–21 (2009).

159. Shumyantseva, V. V, Bulko, T. V & Archakov, A. I. Electrochemical reduction ofcytochrome P450 as an approach to the construction of biosensors and bioreactors.Journal of inorganic biochemistry 99, 1051–63 (2005).

160. Udit, A. K. & Gray, H. B. Electrochemistry of heme-thiolate proteins. Biochemicaland biophysical research communications 338, 470–6 (2005).

161. Hara, M. Application of P450s for biosensing: combination of biotechnology andelectrochemistry. Materials Science and Engineering: C 12, 103–109 (2000).

162. Todorovic, S., Jung, C., Hildebrandt, P. & Murgida, D. H. Conformational transi-tions and redox potential shifts of cytochrome P450 induced by immobilization.Journal of biological inorganic chemistry, JBIC: a publication of the Society of Bio-logical Inorganic Chemistry 11, 119–27 (2006).

97

6. REFERENCES

163. Udit, A. K. et al. Spectroscopy and electrochemistry of cytochrome P450 BM3-surfactant film assemblies. Journal of the American Chemical Society 128, 10320–5(2006).

164. Bonifacio, A. et al. Active-site structure, binding and redox activity of the heme-thiolate enzyme CYP2D6 immobilized on coated Ag electrodes: a surface-enhancedresonance Raman scattering study. Journal of biological inorganic chemistry,JBIC:a publication of the Society of Biological Inorganic Chemistry 13, 85–96 (2008).

165. Schneider, E. & Clark, D. S. Cytochrome P450 (CYP) enzymes and the developmentof CYP biosensors. Biosensors & bioelectronics 39, 1–13 (2013).

166. Alexander I. Archakov & Bachmanova, G. I. Cytochrome P-450 and active oxygen.(1990).

167. Lewis, D. F. V. Guide to Cytochromes P450: Structure and Function. (InformaHealthcare: 2001).

168. Iyanagi, T. Structure and function of NADPH-cytochrome P450 reductase and nitricoxide synthase reductase domain. Biochemical and biophysical research communi-cations 338, 520–8 (2005).

169. Ortiz de Montellano, P. R. Heme Oxygenase Mechanism: Evidence for an Elec-trophilic, Ferric Peroxide Species. Accounts of Chemical Research 31, 543–549 (1998).

170. Nelson, D. R. et al. P450 superfamily: update on new sequences, gene mapping,accession numbers and nomenclature. Pharmacogenetics 6, 1–42 (1996).

171. Im, S.-C. & Waskell, L. The interaction of microsomal cytochrome P450 2B4 withits redox partners, cytochrome P450 reductase and cytochrome b(5). Archives ofbiochemistry and biophysics 507, 144–53 (2011).

172. Ellis, J. et al. Domain motion in cytochrome P450 reductase: conformational equi-libria revealed by NMR and small-angle x-ray scattering. The Journal of biologicalchemistry 284, 36628–37 (2009).

173. Hannemann, F., Bichet, A., Ewen, K. M. & Bernhardt, R. Cytochrome P450 systems–biological variations of electron transport chains. Biochimica et biophysica acta1770, 330–44 (2007).

174. Walker, F. A. & Murataliev, M. B. Electron transfer by diflavin reductases. 1698,1–26 (2004).

175. Smith, G. C., Tew, D. G. & Wolf, C. R. Dissection of NADPH-cytochrome P450 oxi-doreductase into distinct functional domains. Proceedings of the National Academyof Sciences of the United States of America 91, 8710–4 (1994).

98

6. REFERENCES

176. Brenner, S. et al. Inter-flavin electron transfer in cytochrome P450 reductase - effectsof solvent and pH identify hidden complexity in mechanism. The FEBS journal 275,4540–57 (2008).

177. Aigrain, L., Pompon, D., Moréra, S. & Truan, G. Structure of the open conformationof a functional chimeric NADPH cytochrome P450 reductase. EMBO reports 10,742–7 (2009).

178. Iyanagi, T., Makino, N. & Mason, H. S. Redox properties of the reduced nicoti-namide adenine dinucleotide phosphate-cytochrome P-450 and reduced nicotinamideadenine dinucleotide-cytochrome b5 reductases. Biochemistry 13, 1701–10 (1974).

179. Vermilion, J. L. & Coon, M. J. Purified liver microsomal NADPH-cytochrome P-450reductase. Spectral characterization of oxidation-reduction states. The Journal ofbiological chemistry 253, 2694–704 (1978).

180. Zhou, S.-F., Yang, L.-P., Zhou, Z.-W., Liu, Y.-H. & Chan, E. Insights into the substratespecificity, inhibitors, regulation, and polymorphisms and the clinical impact ofhuman cytochrome P450 1A2. The AAPS journal 11, 481–94 (2009).

181. Shumyantseva, V. V., Suprun, E. V., Bulko, T. V., Dobrynina, O. V. & Archakov, a. I.Sensor systems for medical application based on hemoproteins and nanocompositematerials. Biochemistry (Moscow) Supplement Series B: Biomedical Chemistry 4,25–36 (2010).

182. Wilks, A. Analysis of Heme and Hemoproteins: Methods and Protocols. Heme,Chlorophyll, and Bilins (2002).doi:10.1385/1-59259-243-0:157

183. Gómez-Bombarelli, R. et al. Alkylating Potential of Oxetanes. Chemical Research inToxicology 23, 1275–1281 (2010).

184. Kaplan, I. H., Dagcı, K. & Alanyalıoglu, M. Nucleation and Growth Mechanismof Electropolymerization of Methylene Blue: The Effect of Preparation Potential onPoly(methylene blue) Structure. Electroanalysis 22, 2694–2701 (2010).

185. Ni, H., Du, Y., Ma, G., Nagai, M. & Omi, S. Mechanism of Soap-Free EmulsionPolymerization of Styrene and 4-Vinylpyridine: Characteristics of Reaction in theMonomer Phase, Aqueous Phase, and Their Interface. Macromolecules 34, 6577–6585 (2001).

186. Mie, Y., Suzuki, M. & Komatsu, Y. Electrochemically Driven Drug Metabolism byMembranes Containing Human Cytochrome P450. Journal of the American Chem-ical Society 131, 6646–6647 (2009).

187. Estavillo, C. Epoxidation of styrene by human cyt P450 1A2 by thin film electrolysisand peroxide activation compared to solution reactions. Biophysical Chemistry 104,291–296 (2003).

99

6. REFERENCES

188. Bayachou, M., Lin, R., Cho, W. & Farmer, P. J. Electrochemical Reduction of NO byMyoglobin in Surfactant Film: Characterization and Reactivity of the Nitroxyl (NO- ) Adduct. Journal of the American Chemical Society 120, 9888–9893 (1998).

189. Chen, X., Hu, N., Zeng, Y., Rusling, J. F. & Yang, J. Ordered Electrochemically ActiveFilms of Hemoglobin, Didodecyldimethylammonium Ions, and Clay. Langmuir 15,7022–7030 (1999).

190. Matsumura, H., Nakamura, N., Yohda, M. & Ohno, H. The electrochemical proper-ties of thermophilic cytochrome P450 CYP119A2 at extremely high temperatures inpoly(ethylene oxide). Electrochemistry Communications 9, 361–364 (2007).

191. Krishnan, S., Schenkman, J. B. & Rusling, J. F. Bioelectronic Delivery of Electronsto Cytochrome P450 Enzymes. The Journal of Physical Chemistry B 0, 8371–8380(2011).

192. Bartlett, P. N. & Techniques, E. Bioelectrochemistry: fundamentals, experimentaltechniques and applications. Experimental Techniques (John Wiley & Sons: Chich-ester, England; Hoboken, NJ, 2008).

193. Nalwa, E. H. S., Bauza, D., Derry, G. N., Miller, R. & Fainerman, V. B. Handbookof Surfaces and Interfaces of Materials. Handbook of Surfaces and Interfaces ofMaterials 1, xxv–xxviii (2001).

194. Limoges, B. & Savéant, J.-M. Catalysis by immobilized redox enzymes. Diagnosisof inactivation and reactivation effects through odd cyclic voltammetric responses.Journal of Electroanalytical Chemistry 562, 43–52 (2004).

195. Gwyer, J. D., Angove, H. C., Richardson, D. J. & Butt, J. N. Redox-triggered eventsin cytochrome c nitrite reductase. Bioelectrochemistry (Amsterdam, Netherlands)63, 43–7 (2004).

196. Dawson, J. Probing structure-function relations in heme-containing oxygenases andperoxidases. Science 240, 433–439 (1988).

197. Liu, S. & Ju, H. Reagentless glucose biosensor based on direct electron transfer ofglucose oxidase immobilized on colloidal gold modified carbon paste electrode.Biosensors and Bioelectronics 19, 177–183 (2003).

198. Boccaccini, A. R. et al. Electrophoretic deposition of carbon nanotubes. Carbon 44,3149–3160 (2006).

199. Cho, J. et al. Characterisation of carbon nanotube films deposited by electrophoreticdeposition. Carbon 47, 58–67 (2009).

100

6. REFERENCES

200. Goodwin, J. W., Hearn, J., Ho, C. C. & Ottewill, R. H. Studies on the preparation andcharacterisation of monodisperse polystyrene laticee. Colloid and Polymer ScienceKolloid Zeitschrift & Zeitschrift for Polymere 252, 464–471 (1974).

201. Mazurenko, I. et al. Electrophoretic deposition of macroporous carbon nanotubeassemblies for electrochemical applications. Carbon 53, 302–312 (2013).

202. Plumeré, N., Henig, J. & Campbell, W. H. Enzyme-catalyzed O2 removal system forelectrochemical analysis under ambient air: application in an amperometric nitratebiosensor. Analytical chemistry 84, 2141–6 (2012).

203. Alayash, A. I., Patel, R. P. & Cashon, R. E. Redox reactions of hemoglobin and myo-globin: biological and toxicological implications. Antioxidants & redox signaling 3,313–27 (2001).

204. Nagababu, E. & Rifkind, J. M. Heme degradation during autoxidation of oxyhe-moglobin. Biochemical and biophysical research communications 273, 839–45 (2000).

205. Nagababu, E. & Rifkind, J. M. Reaction of hydrogen peroxide with ferrylhemoglobin:superoxide production and heme degradation. Biochemistry 39, 12503–11 (2000).

206. Nagababu, E., Ramasamy, S., Rifkind, J. M., Jia, Y. & Alayash, A. I. Site-specificcross-linking of human and bovine hemoglobins differentially alters oxygen bind-ing and redox side reactions producing rhombic heme and heme degradation. Bio-chemistry 41, 7407–15 (2002).

207. Almeida, M. G. et al. A needle in a haystack: the active site of the membrane-boundcomplex cytochrome c nitrite reductase. FEBS letters 581, 284–8 (2007).

208. Villegas, J. a, Mauk, a G. & Vazquez-Duhalt, R. A cytochrome c variant resistant toheme degradation by hydrogen peroxide. Chemistry & biology 7, 237–44 (2000).

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